Microbial nitrous oxide production and nitrogen cycling ... · Nitrogen cycling is intimately...

Microbial nitrous oxide production and nitrogen cycling associated with aquatic invertebrates

Dissertation

zur Erlangung des Doktorgrades

der Naturwissenschaften

-Dr.rer.nat.-

dem Fachbereich Biologie/Chemie

der Universität Bremen

vorgelegt von

Ines Heisterkamp

Bremen

Juni 2012

Die vorliegende Arbeit wurde in der Zeit von Januar 2009 bis Juni 2012 am

Max-Planck-Institut für marine Mikrobiologie in Bremen angefertigt.

1. Gutachter: Prof. Dr. Bo Barker Jørgensen

2. Gutachter: Prof. Dr. Ulrich Fischer

Prüfer:

Dr. Peter Stief

Prof. Dr. Victor Smetacek

Tag des Promotionskolloquiums: 6. August 2012

2

Table of contents

Table of contents

Summary .........................................................................................................................5

Zusammenfassung ..........................................................................................................7

Chapter 1 .......................................................................................................................11

General introduction ........................................................................................12

Aims of the thesis ..............................................................................................44

Overview of manuscripts .................................................................................46

References .........................................................................................................49

Chapter 2 .......................................................................................................................63

Nitrous oxide production associated with coastal marine invertebrates

Chapter 3 .......................................................................................................................85

Shell biofilm-associated nitrous oxide production in marine molluscs:

processes, precursors and relative importance

Chapter 4 .....................................................................................................................117

Shell biofilm nitrification and gut denitrification contribute to emission of

nitrous oxide by the invasive freshwater mussel Dreissena polymorpha

(Zebra Mussel)

Chapter 5 .....................................................................................................................133

Incomplete denitrification in the gut of the aquacultured shrimp

Litopenaeus vannamei as source of nitrous oxide

Chapter 6 .....................................................................................................................155

Indirect control of the intracellular nitrate pool of intertidal sediment by

the polychaete Hediste diversicolor

Chapter 7 .....................................................................................................................183

Conclusion and perspectives

Contributed works......................................................................................................203

Danksagung.................................................................................................................211

3

Summary

Summary

Nitrogen cycling is intimately linked to the activity of microorganisms that mediate the

diverse nitrogen transformations and play a fundamental role in regulating the fate of

nitrogen in the Earth’s terrestrial and aquatic ecosystems. Microbial activity is

influenced by physical, chemical, and biological factors that can be profoundly shaped

by macrofaunal organisms, especially in benthic aquatic systems. This thesis therefore

aimed at investigating the interactions between microorganisms and benthic aquatic

invertebrates and their role in biogeochemical nitrogen cycling, especially regarding the

production of nitrous oxide (N2O). This intermediate and by-product of microbial

nitrogen cycling processes (mainly nitrification and denitrification) is of great

importance as a greenhouse gas and ozone-depleting substance in the atmosphere. To

date, the biogenic N2O sources remain poorly quantified in the global N2O budget.

Natural N2O production mainly takes place in soils, sediments, and water bodies, but

also occurs in the anoxic gut of earthworms and freshwater invertebrates.

This thesis investigated for the first time the N2O emission potential of marine

invertebrates that densely colonize coastal benthic ecosystems. An initial screening

effort in the German Wadden Sea and Aarhus Bay, Denmark, revealed a large variety of

marine invertebrate species as N2O emitters (Chapter 2). Statistical analysis showed that

the N2O emission potential is not restricted to a certain taxonomic group or feeding

guild, but rather correlates with body weight, habitat, and the presence of microbial

biofilms on the shell or exoskeleton of the animals. This suggests that N2O emission

from marine invertebrates is not necessarily due to denitrification in the gut, but may

also result from microbial activity on the external surfaces of the animal.

The novel pathway of N2O production in shell biofilms was investigated in detail by a

combination of short-term and long-term incubation experiments, stable isotope

experiments, microsensor measurements, and molecular analysis (Chapters 3 and 4).

Investigations on three marine (Mytilus edulis, Littorina littorea, Hinia reticulata) and

one freshwater mollusc species (Dreissena polymorpha) revealed that shell biofilms

significantly contribute to the total animal-associated N2O production via both

denitrification and nitrification. Ammonium excretion by the molluscs was sufficient to

5

Summary

sustain nitrification-derived N2O production in the shell biofilms and thus potentially

decouples invertebrate-associated N2O production from environmental nitrogen

concentrations. This was demonstrated in detail for the snail H. reticulata, which

promotes growth and N2O production of its shell biofilm by enriching its immediate

surroundings with dissolved inorganic nitrogen.

The shrimp Litopenaeus vannamei, the most important crustacean species in

aquaculture worldwide, was found to emit N2O at the highest rate recorded for any

marine invertebrate so far (Chapter 5). The shrimp gut represents a transient anoxic

habitat in which ingested bacteria produce N2O due to incomplete denitrification. At

high stocking densities, L. vannamei may significantly contribute to the N2O

supersaturation observed in the rearing tank of the shrimp aquaculture.

In an additional study, the fate of nitrogen was investigated in an animal-bacteria-

microalgae interaction occurring in intertidal flats (Chapter 6). Diatoms were found to

store more nitrate intracellularly when the polychaete Hediste diversicolor stimulated

the activity of nitrifying bacteria by excretion of ammonium and oxygenation of the

sediment. This intricate interplay alters the forms and availability of the important

nutrient nitrogen in marine sediments.

Conceptually, benthic invertebrates represent “hotspots” of microbial nitrogen cycling

that add specific features to the general marine nitrogen cycle, such as the noticeable

N2O production and the partial decoupling of microbial activity from ambient nutrient

supply. In particular, this thesis revealed that invertebrate-associated N2O production

constitutes an important link between reactive nitrogen in aquatic environments and

atmospheric N2O and is controlled by environmental, autecological, and physiological

factors.

6

Zusammenfassung

Zusammenfassung

Der Stickstoffkreislauf ist aufs Engste mit der Aktivität von Mikroorganismen

verknüpft, die verschiedenste Stickstoffumwandlungen durchführen und eine

entscheidende Rolle für die Umsetzung sowie den Verbleib von Stickstoffverbindungen

in terrestrischen und aquatischen Ökosystemen spielen. Die mikrobielle Aktivität wird

durch verschiedene physikalische, chemische und biologische Faktoren reguliert,

welche maßgeblich durch wirbellose Tiere (Invertebraten) beeinflusst werden können.

Dies gilt in besonderer Weise für das Benthos aquatischer Ökosysteme. Das Ziel der

vorliegenden Arbeit war es daher, die Interaktionen zwischen Mikroorganismen und

benthischen Invertebraten sowie ihre Rolle im biogeochemischen Stickstoffkreislauf zu

erforschen. Schwerpunktmäßig wurde dabei die Produktion von Distickstoffmonoxid

(N2O), auch bekannt als Lachgas, untersucht. Dieses Zwischen- und Nebenprodukt

zahlreicher mikrobieller Stickstoffumwandlungen (hauptsächlich Nitrifikation und

Denitrifikation) ist von globaler Bedeutung, da es in der Erdatmosphäre signifikant zum

Treibhauseffekt und zum Ozonabbau beiträgt. Im globalen N2O-Budget sind die

biogenen Quellen von N2O allerdings bis heute nur unvollständig quantifiziert. Biogene

Produktion von N2O findet hauptsächlich im Boden sowie in den Sedimenten und der

Wassersäule aquatischer Ökosysteme statt, wurde aber auch in den sauerstofffreien

Därmen von Regenwürmern und Süßwasser-Invertebraten beobachtet.

In der vorliegenden Arbeit wurde nun zum ersten Mal das N2O-Emissionspotenzial

mariner Invertebraten untersucht, die küstennahe Sedimente dicht besiedeln. Zu Beginn

der Arbeit wurde ein Screening verschiedener Tierarten aus dem deutschen Wattenmeer

und der Bucht von Aarhus in Dänemark durchgeführt (Kapitel 2). Dabei erwiesen sich

zahlreiche marine Invertebraten-Arten als N2O-Emittenten. Eine statistische Analyse

zeigte, dass ein vorliegendes N2O-Emissionspotenzial nicht auf bestimmte

taxonomische Gruppen und Ernährungstypen beschränkt ist, sondern mit dem

Körpergewicht, dem Habitat und dem Vorhandensein von mikrobiellen Biofilmen auf

der Schale oder dem Exoskelett der Tierarten korreliert. Diese Befunde deuteten

erstmals darauf hin, dass die N2O-Emission mariner Invertebraten nicht zwangsläufig

durch Denitrifikation im Darm bedingt ist, sondern auch auf mikrobielle Aktivitäten auf

der Oberfläche des Tieres zurückgehen kann.

7

Zusammenfassung

Dieser neue N2O-Produktionsweg in „Schalen-Biofilmen“ wurde im Detail mit

Kurzzeit- und Langzeit-Inkubationsexperimenten, stabilen Isotopen-Experimenten,

Mikrosensormessungen und molekularbiologischen Analysen untersucht (Kapitel 3 und

4). Untersuchungen von drei marinen (Mytilus edulis, Littorina littorea, Hinia reticulata)

und einer Süßwasser-Molluskenart (Dreissena polymorpha) ergaben, dass „Schalen-

Biofilme“ signifikant zur N2O-Emission des gesamten Tieres beitragen, und zwar

sowohl aufgrund von Denitrifikation als auch Nitrifikation. Die Exkretion von

Ammonium durch die Mollusken war stets ausreichend, um die N2O-Produktion durch

Nitrifikation aufrechtzuerhalten, und kann demnach die mit dem Tier assoziierte N2O-

Emission von der Stickstoffkonzentration im umgebenden Wasser entkoppeln. Dieses

wurde im Detail für die Schneckenart H. reticulata nachgewiesen, welche das

Wachstum und die N2O-Produktion von „Schalen-Biofilmen“ fördert, indem sie ihre

eigene unmittelbare Umgebung mit gelöstem inorganischem Stickstoff anreichert.

Die Garnele Litopenaeus vannamei ist die weltweit wichtigste Crustaceen-Art in

Aquakultur und emittiert N2O mit der höchsten Rate, die bislang für marine

Invertebraten gemessen werden konnte (Kapitel 5). Der Darm der Garnele stellt für

ingestierte Bakterien ein anoxisches Kurzzeithabitat dar, in dem sie N2O durch

unvollständige Denitrifikation produzieren. Bei hoher Besatzdichte trägt L. vannamei

vermutlich signifikant zu der in den Zuchtbecken beobachteten N2O-Übersättigung bei.

In einer weiteren Studie wurde untersucht, wie eine Tier-Mikroben-Mikroalgen

Interaktion den Stickstoffkreislauf in Wattsedimenten beeinflusst (Kapitel 6). Es zeigte

sich, dass Diatomeen dann mehr Nitrat intrazellulär speichern, wenn die Polychaeten-

Art Hediste diversicolor die Aktivität nitrifizierender Bakterien durch Exkretion von

Ammonium und Oxygenierung des Sedimentes stimuliert. Dieses komplexe

Zusammenspiel führt dazu, dass Form und Verfügbarkeit von Stickstoff als überaus

wichtigem Nährstoff in marinen Sedimenten verändert werden.

Konzeptionell stellen benthische Invertebraten „hotspots“ mikrobieller Stickstoff-

umsetzungen dar, die den allgemeinen Stickstoffkreislauf mit besonderen Leistungen

ergänzen, wie z.B. mit einer beachtlichen N2O-Produktion und einer teilweisen

Entkopplung mikrobieller Aktivität von der Nährstoffzufuhr in der Umwelt.

Insbesondere konnte in der vorliegenden Arbeit gezeigt werden, dass die Invertebraten-

8

Zusammenfassung

assoziierte N2O-Produktion eine wichtige Verbindung zwischen reaktiven

Stickstoffverbindungen in aquatischen Ökosystemen und dem atmosphärischen N2O

darstellt, die maßgeblich durch autökologische, physiologische und Umweltfaktoren

bestimmt wird.

9

Chapter 1

11

Chapter 1 General introduction

General introduction

This thesis aimed at investigating interactions between microorganisms and benthic

aquatic invertebrates and their role in biogeochemical nitrogen cycling. The primary

objective of this thesis was to unravel the potential and the underlying mechanisms of

microbial N2O production associated with aquatic invertebrates (Chapters 2 to 5). In

addition, the impact of an invertebrate-bacteria-microalgae interaction on the intra-

cellular nitrate pool in marine sediment was investigated (Chapter 6). To give a

background for the following Chapters the general introduction will give an overview

on 1) the major processes of the nitrogen cycle and the role of nitrogen as important

nutrient, 2) the effects of nitrous oxide, its sources and sinks, and the estimated global

N2O-budget, 3) the pathways of microbial N2O production and their controlling factors,

4) nitrogen cycling and important sites and processes of N2O production in aquatic

environments, 5) the effects of invertebrates on nitrogen turnover in aquatic

environments, and finally on 6) N2O emission by benthic macrofauna and other

organisms.

The nitrogen cycle – processes, environmental importance and anthropogenic alteration

Nitrogen (N) is a key element for life on Earth, as all living organisms require nitrogen

for the synthesis of proteins, nucleic acids, and other important N-containing

biomolecules. It exists in a multiplicity of organic and inorganic forms and in a wide

range of oxidation states, ranging from −III in ammonium (NH4+) and organic matter to

+V in nitrate (NO3−) (Hulth et al. 2005, Gruber 2008). The N cycle is almost entirely

dependent on redox reactions (Figure 1). These chemical transformations are primarily

mediated by microorganisms that use nitrogen to synthesize biomass or to gain energy

for growth (Zehr & Ward 2002, Canfield et al. 2010). Microorganisms are therefore key

players in biogeochemical cycling of nitrogen, which mainly takes place in soils,

sediments and water bodies.

12


Processes of the nitrogen cycle

N2-fixing microorganisms play a fundamental role in the nitrogen cycle, since they are

the only organisms that can use the huge reservoir of dinitrogen gas (N2) in the

atmosphere (Carpenter & Capone 2008). With help of their nitrogenase enzyme

complex, they break the very strong triple bond of N2 and reduce N2 to NH4+ that is

incorporated into particulate organic nitrogen (PON). All other organisms rely on the

supply of fixed nitrogen forms, also referred to as reactive nitrogen (Nr).

Microorganisms and plants take up dissolved inorganic nitrogen (DIN = NH4+, nitrite

(NO2−), and NO3

−) from the environment and assimilate it into PON (Oaks 1992,

Mulholland & Lomas 2008). In addition, microorganisms and some plants can use

dissolved organic nitrogen (DON) compounds (e.g., urea and amino acids, Jones et al.

2004, Bradley et al. 2010). Animals meet their nitrogen requirements by feeding on

PON. The organic nitrogen in living and dead organisms is recycled back to inorganic

nitrogen by remineralization processes. Heterotrophic microbes and animals degrade the

N-containing macromolecules and subsequently release NH4+ and DON (Canfield et al.

2005).

In the presence of oxygen (O2), NH4+ is oxidized over NO2

− to NO3− by chemo-

lithotrophic bacteria and archaea in a process known as nitrification (Ward 2008). The

gas nitrous oxide (N2O) is produced as a by-product in this process. The resulting

oxidized compounds NO2− and NO3

− (NOx−) are used as electron acceptors by diverse

groups of microorganisms when the terminal electron acceptor O2 is limiting. NOx− is

either reduced to NH4+ by a process called dissimilatory nitrate reduction to ammonium

(DNRA) or to N2 by the process of denitrification (Lam & Kuypers 2011). Both

processes are mainly carried out by heterotrophic bacteria, but nitrate reduction can also

be coupled to the oxidation of inorganic compounds by chemolithotrophs.

Denitrification produces N2O as an intermediate (Knowles 1982), whereas DNRA is

thought to produce trace amounts of N2O as by-product (Kelso et al. 1997, Cruz-Garcia

et al. 2007). Besides denitrification, N2 is also produced by anaerobic ammonium

oxidation (Mulder et al. 1995, Strous et al. 1999). During this so-called anammox

process, anaerobic chemoautotrophic bacteria within the group of planctomycetes

produce N2 by coupling the reduction of NO2− with the oxidation of NH4

+. The

13


anammox- and DNRA-bacteria are generally strict anaerobes, while most denitrifying

bacteria are facultative anaerobes. Denitrification and anammox play a fundamental role

in the N cycle by returning N2 gas back to the atmosphere and thus reducing the amount

of biologically available nitrogen (Devol 2008). The N2O produced during nitrogen

cycling is either consumed by denitrification or escapes to the atmosphere.

V

IV

III

II

I

0

-I

-II

-III

Oxid

ation s

tate

NO3−

NH4+

N2

NH2OH

NO2−

NO

Norg

NO2−

N2O

Remineralization

N2O

Figure 1: Major chemical species and transformations of the nitrogen cycle. The various chemical N-species are plotted versus their oxidation state. Major processes involved in N cycling: fixation of N2 to NH4

+ (green); assimilation of NO3−, NO2

−, and NH4+ to organic

nitrogen (Norg) (black); remineralization of Norg to NH4+ (brown); oxidation of NH4

+ to NO3− via

nitrification (red), reduction of NO3− to N2 via denitrification (blue); dissimilatory reduction of

NO3− to NH4

+ (DNRA, purple); and anaerobic oxidation of NH4+ to N2 (anammox, orange). N2O

is produced as by-product during nitrification and DNRA (dashed lines) and as intermediate in denitrification.

Environmental importance and anthropogenic alteration of the N cycle

The nitrogen cycle is of particular interest, as the availability of nitrogen influences the

rate of key processes in terrestrial and aquatic ecosystems, such as primary production

and decomposition of organic matter. It thereby interacts with biogeochemical cycles of

many other elements, in particular carbon (Gruber & Galloway 2008). The scarcity of

fixed inorganic nitrogen limits primary production in many marine and terrestrial

ecosystems (Falkowski 1997, Vitousek et al. 2002). The nitrogen availability

14


consequently controls the amount of carbon dioxide (CO2) that is fixed by plants or

phytoplankton and thereby strongly influences the short-term sequestration of the

greenhouse gas CO2 in terrestrial ecosystems and the deep oceans (Falkowski et al.

1998, Zaehle et al. 2011). The global N cycle is thus fundamental to the functioning of

the Earth’s climate (Vitousek et al. 1997, Holland et al. 2005).

Over the last century, human activities have substantially altered the global N cycle by

tremendously increasing the amount of reactive nitrogen in the biosphere (Galloway et

al. 2008). The increase in Nr is largely due to two anthropogenic activities: (i) food

production promoted by application of synthetic fertilizers and cultivation of N2-fixing

crops, and (ii) energy production by fossil fuel combustion (Vitousek et al. 1997,

Galloway et al. 2004). Anthropogenic Nr sources provide nowadays almost 50% of the

total fixed nitrogen produced annually on Earth (Canfield et al. 2010 and references

therein). A significant fraction of the Nr applied on agricultural soils leaks into rivers,

lakes, and aquifers, and is transported to coastal ecosystems (Boyer et al. 2006,

Schlesinger 2009, Seitzinger et al. 2010), or evaporates as NH3 or NOx (NO + NO2) and

is globally distributed through atmospheric transport and subsequent deposition

(Galloway & Cowling 2002). Anthropogenic N thus influences biogeochemical

processes in terrestrial, freshwater, coastal, and oceanic ecosystems (Galloway et al.

2004, Duce et al. 2008).

The anthropogenic perturbation of the N cycle causes substantial and manifold

environmental problems (Vitousek et al. 1997, Matson et al. 2002, Rabalais 2002).

Among these are eutrophication of terrestrial and aquatic systems, acidification of soils

and freshwaters, and increased emission of the greenhouse gas nitrous oxide. The

massive acceleration of the N cycle is projected to further increase to sufficiently meet

the human dietary and energy demands of a growing world population (Galloway et al.

2008). Efficient management in food and energy production and improved

understanding of mechanisms controlling the fate of Nr in the environment are urgently

needed to reduce the adverse effects of Nr on the Earth’s biosphere and climate. This

especially includes the pathways and mechanisms leading to the emission of the

greenhouse gas N2O, which are to date not satisfactorily understood (Galloway et al.

2008, Davidson 2009, Butterbach-Bahl & Dannenmann 2011).

15


Nitrous oxide – its properties, sources and sinks

Properties of nitrous oxide

Nitrous oxide (dinitrogen monoxide), also known as laughing gas, is a natural

atmospheric trace gas with the molecular formula N2O and a molar mass of 44 g mol−1.

Its solubility in water is very high with 24.1 mmol L−1 at a salinity of 34 and

temperature of 20°C (Weiss & Price 1980). The present atmospheric concentration of

N2O is far higher than at any time during the past 140,000 years (Schilt et al. 2010).

Within the past 10,000 years, changes in atmospheric N2O concentration were relatively

small until the beginning of the industrial era (Figure 2, Forster et al. 2007). Since then

the atmospheric N2O concentration has increased tremendously from 270 ppb in 1750 to

320 ppb in 2005. Within the last few decades, the concentration increased with a rate of

0.2−0.3% per year (Forster et al. 2007).

Figure 2: Atmospheric concentrations of nitrous oxide over the last 10,000 years (large panel) and since 1750 (inset panel). Measurements are shown from ice cores (symbols with different colours for different studies) and atmospheric samples (red lines). The corresponding radiative forcing is shown on the right hand axis of the large panel. The radiative forcing is a measure of the influence a factor has in altering the balance of incoming and outgoing energy in the Earth-atmosphere system and is an index of the importance of the factor as a potential climate change mechanism. A positive forcing (more incoming energy) tends to warm the system. Figure and text are taken from the Fourth Assessment Report of the Intergovernmental Panel on Climate Change (IPCC 2007).

16


The increasing atmospheric N2O concentration is of particular interest, since N2O

greatly impacts the chemistry of the Earth’s troposphere and stratosphere (Figure 3). In

the troposphere, N2O acts as a strong greenhouse gas by absorbing and re-emitting part

of the infrared radiation coming from the Earth’s surface and thereby heating the Earth

system (Forster et al. 2007, Wuebbles 2009). Per molecule N2O has an approximately

300 times higher global warming potential over a 100-year timescale than CO2 and a

particularly long atmospheric lifetime of about 120 years (Forster et al. 2007). It

accounts for approximately 7-10% of the overall anthropogenic greenhouse effect and is

the third most important human-induced greenhouse gas after CO2 and methane (CH4)

(IPCC 2007).

In addition to its global warming effect in the troposphere, N2O also plays a key role in

the destruction of the stratospheric ozone layer (Ravishankara et al. 2009). Since N2O is

not removed from the troposphere by chemical reactions, it reaches the stratosphere

where it reacts with excited oxygen (O(1D)) to form nitric oxide (NO) (Schlesinger

1997, Olsen et al. 2001). NO in turn reacts with ozone (O3) to form nitrogen dioxide

(NO2). The produced nitrogen oxides (NO + NO2) destroy ozone via following reactions

NO + O3 � NO2 + O2

NO2 + O(1D) � NO + O2 (Crutzen 1970, Johnston 1971).

Nearly all stratospheric NO is produced from N2O, which is currently the most

important ozone-depleting substance and is expected to remain the largest one

throughout the 21st century (Ravishankara et al. 2009).

17


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��

��

��

��

��

��

��

Figure 3: The impact of N2O on the greenhouse effect and the destruction of the ozone layer. Solar radiation passes through the atmosphere, some radiation is reflected back, the remainder is absorbed by the Earth’s surface. Here, it is converted into heat causing the emission of long-wave (infrared) radiation back to the atmosphere. In the troposphere, N2O and other greenhouse gases absorb part of the infrared radiation and re-emit it back to the Earth’s surface, thus warming up the Earth and the troposphere. The reflected radiation is re-emitted from the Earth’s surface and is lost in space when it passes through the troposphere. In the stratosphere, N2O is oxidized to NO or photolysed to N2 and O(1D) species. NO and O(1D) react with O3 and destroy the ozone layer.

Sources and sinks of nitrous oxide

A wide range of N2O sources of natural and anthropogenic origin have been identified

within the last few decades. However, the uncertainty ranges of the individual sources

are high and there still remain unknown sites, mechanisms of production and regulating

factors that need to be identified to refine the global N2O budget (Forster et al. 2007,

Rubasinghege et al. 2011). Syakila & Kroeze (2011) present the most recent estimates

on the global N2O budget and calculated the global N2O emission to be 18.3 Tg N yr−1

in the year 2000 (Table 1). Global N2O emissions thus increased compared to the

estimates for the 1990s (Table 1, Denman et al. 2007). Natural sources are estimated to

account for 60% and anthropogenic sources for 40% of the global N2O emissions

18


(Denman et al. 2007, Syakila & Kroeze 2011). Natural N2O emissions derive primarily

from soils (~ 60%) and oceans (~ 35%) via microbial N conversion processes (Denman

et al 2007, Syakila & Kroeze 2011). Agriculture is the most important anthropogenic

source and is responsible for 60−70% of the anthropogenic N2O emissions (recent

detailed inventories of agricultural emissions are available in Davidson 2009). The

remaining 30−40% of the anthropogenic N2O emissions arise from fossil fuel

combustion, industrial processes, biomass and biofuel burning (Crutzen et al. 2008).

Agricultural activities do not only lead to direct N2O emissions from fertilized soils and

from animal production, but also to indirect N2O emissions when fixed nitrogen applied

to agricultural systems is released to natural environments by leaching, sewage, and

atmospheric deposition of nitrogen oxides and ammonia (NH3) (Mosier et al. 1998).

Enhanced N2O production in rivers, estuaries and coastal zones due to anthropogenic N

input represent an important source of N2O and are estimated to be 1.7 Tg N yr−1

(Denman et al. 2007). However, the estimates for N2O emissions from aquatic

ecosystems remain highly uncertain (Nevison et al. 2004, Baulch et al. 2011). Beaulieu

and coworkers (2010), for instance, calculated N2O emission from river networks to be

0.68 Tg N yr−1, which is three times higher than the emissions estimated by Denman et

al. 2007. Seitzinger et al. (2000) calculated that rivers, estuaries, and continental shelves

make up 35% of the total aquatic N2O emissions and the open oceans the remaining

65%, while Bange et al. (1996) proposed that estuaries and continental shelves

contribute as much as 60% to the total oceanic N2O emission. Since anthropogenic N

even reaches the open oceans and increases oceanic N2O emission (Duce et al. 2008,

Suntharalingam et al. 2012), oceans are included as both natural and anthropogenic

sources in the updated N2O budget (Table 1, Syakila & Kroeze 2011). Another source

that is not yet included in the global N2O budget is the N2O emission from aquaculture.

William and Crutzen (2010) estimated that N2O emission from aquaculture accounts for

0.12 Tg N yr−1 and argued that this emission is likely to increase to 1.01 Tg N yr−1

within the next 20 years due to the rapid global growth rate of aquaculture industry.

This estimate is solely based on theoretical N2O emissions from nitrogenous waste of

aquacultures. However, in Chapter 5 of this thesis, it is shown that also the aquacultured

animals themselves can emit N2O and thus represent an additional source of N2O

emission from aquacultures.

19


In contrast to the various sources of N2O, the only major sinks of N2O are the oxidation

to NO or the photolysis to N2 and O(1D) in the stratosphere (Khalil et al. 2002). In

addition, denitrification can act as a net sink of N2O in forest soils, some aquatic

systems, and riparian zones (reviewed in Chapuis-Lardy et al. 2007, Billings 2008).

However, this surface uptake of N2O is estimated to be about 0.01 Tg N yr−1 and thus of

minor importance for the global N2O budget (Syakila & Kroeze 2011).

Table 1: Estimates of the global N2O budget for the 1990s from the Fourth Assessment Report of the Intergovernmental Panel on Climate Change (IPCC, Denman et al 2007) and newest available estimates from Syakila & Kroeze (2011) for the year 2000 (Tg N yr−1). Ranges of estimates are presented in brackets.

1990s 2000

Sources of N2O

Natural 11 10.5

Soil 6.6 (3.3−9.0) 6−7

Ocean 3.8 (1.8−5.8) 3−4

Atmospheric chemistry 0.6 (0.3−1.2) <1

Anthropogenic 6.7 7.8

Energy, industry, biomass burning 2.0 (0.7−3.7)a 1.9

Agriculture (including animal production) 4.7 (2.3−8.0) 4.9

Direct emissions 2.8 (1.7−4.8) 3.8

Indirect emissions 1.9 (0.6−3.2) 1.1

Human excreta/sewage 0.2 (0.1−0.3) −

Rivers, estuaries and coastal zones 1.7 (0.5−2.9) −

Oceans − 1.0

Total 17.7 (8.5−27.7) 18.3

Sinks of N2O

Surface sink − 0.01

Stratospheric sink 12.5 (10.0−15.0)

Net atmospheric increase 6.8a Including N2O from atmospheric deposition, which is in part agricultural

20


Pathways of microbial nitrous oxide production

Biogenic N2O production in natural and anthropogenically influenced ecosystems

originates primarily from microbial processes. Although a wide range of microbial

pathways has the potential to produce N2O, its production in soils, sediments and water

bodies is mainly ascribed to two processes: denitrification and nitrification (Bange 2008,

Kool et al. 2011).

Denitrification

Denitrification, the respiratory reduction of NO3− or NO2

− to N2, is typically considered

to be the dominant N2O source in soils, sediments and anoxic water bodies, as it is

induced under low O2 or anoxic conditions (Codispoti et al. 2001, Bouwman et al.

2002). This facultative anaerobic respiration process is phylogenetically widespread,

occurring in the domains Bacteria (Zumft 1997), Archaea (Cabello et al. 2004), and few

Eukaryota such as fungi (Shoun et al. 1992) and foraminifera (Risgaard-Petersen et al.

2006). Most research has focused on denitrifiers within the proteobacteria (alpha, beta,

gamma, and epsilon divisions), since these are generally believed to be the dominant

denitrifying organisms in most environments (Wallenstein et al. 2006).

The complete denitrification pathway (NO3− � NO2

− � NO � N2O � N2) involves

four enzymatically catalyzed reduction steps (Figure 4 and 6). In bacteria, the

dissimilatory reduction of NO3− to NO2

− is mediated either by the membrane-bound

nitrate reductase (NAR) that has its active site in the cytoplasm or by the periplasmic

nitrate reductase (NAP) (Gonzalez et al. 2006). These nitrate reductases are, however,

also present in nitrate-reducing bacteria that do not denitrify (e.g., DNRA bacteria)

(Zumft 1997, Wallenstein et al. 2006, Richardson et al. 2009). The NO2− produced is

then reduced to NO by a periplasmic nitrite reductase (NIR). Two evolutionarily

unrelated forms of the NIR enzyme exist, a copper-containing reductase, encoded by the

nirK gene, and a cytochrome cd1-nitrite reductase, encoded by the nirS gene (Zumft

1997). Reduction of the highly reactive and toxic NO to the non-toxic N2O is catalyzed

by the nitric oxide reductase (NOR), an integral membrane protein with its active site in

the periplasm. NIR and NOR are controlled interdependently at both the transcriptional

21


and enzyme level to prevent accumulation of NO (Ferguson 1994, Zumft 1997). The

periplasmic nitrous oxide reductase (NOS) mediates the reduction of N2O to N2 and is

thus crucial for determining whether denitrification acts as a source or sink of N2O. The

described anaerobic reaction chain is split over the periplasmic and cytoplasmic

compartments, which allows the formation of a proton gradient across the bacterial

membrane that is used for synthesis of ATP and NADH (Zumft 1997, Kraft et al. 2011).

Figure 4: Organization and sidedness of the anaerobic electron transfer chain of the denitrifying bacterium Pseudomonas stutzeri. The shaded areas represent the components of the constitutive aerobic respiratory chain consisting of an NADH dehydrogenase complex (DH), quinone cycle (Q, QH2), cytochrome bc1 complex (Cyt bc1), and the cytochrome cb terminal oxidase complex (Cyt cb). The respiratory denitrification system comprises membrane-bound (NAR) and periplasmic (NAP) NO3

− reductases, NO2− reductase (NIR), NO reductase (NOR), and N2O

reductase (N2OR). Abbreviations: FeS, iron-sulfur centers; b, c, and d1, heme B, heme C, and heme D1, respectively; cyt c, unspecified c-type cytochromes accepting electrons from the bc1complex and acting on N2OR and NOR; cyt c551, cytochrome c551; AP, postulated NO3

−/ NO2−

antiporter. Figure and text are taken from Zumft (1997).

Denitrification is coupled to the oxidation of organic carbon (Corg) in heterotrophic

denitrifiers or to the oxidation of inorganic compounds such as ferrous iron, reduced

sulfur compounds, and hydrogen in autotrophic denitrifiers (Knowles 1982, Zumft 1997,

Straub & Buchholz-Cleven 1998). Overall, denitrification enzymes are induced when

O2 concentrations are low and oxidized inorganic N compounds as well as appropriate

organic or inorganic electron donors are available (Tiedje 1988). Denitrification can,

however, also occur under oxic conditions, as has been shown for several isolated

bacterial species (Robertson & Kuenen 1984, Robertson et al. 1989, Patureau et al.

22


2000), and for microbial communities in aquatic (Trevors & Starodub 1987, Gao et al.

2010) and terrestrial environments (Lloyd 1993). This aerobic denitrification often

results in the accumulation of N2O as do shifts from anoxic to oxic conditions, since the

NOS enzyme of many denitrifiers shows a higher sensitivity towards O2 than the other

denitrification enzymes (Bonin & Raymond 1990, Frette et al. 1997, Patureau et al.

2000). Furthermore, it is common that denitrifying bacteria do not possess all four

reductases and are consequently not capable to perform the complete denitrification

process. It is estimated that the percentage of N2O-respiring taxa is only 10−15% of all

known denitrifying taxa (Zumft & Kroneck 2007). Therefore, N2O is also primarily

produced when bacterial strains containing NOS are underrepresented in the bacterial

community (Zumft 1997, Gregory et al. 2003).

Nitrification

Nitrification is the stepwise aerobic oxidation of NH4+ to NO2

− and further to NO3−. The

oxidation of NH4+ to NO2

− is performed by chemolithoautotrophic ammonia-oxidizing

bacteria (AOB) and, as recently discovered, by ammonia-oxidizing archaea (AOA)

(Kowalchuk & Stephen 2001, Konneke et al. 2005). The second step, the oxidation of

NO2− to NO3

−, is mediated by a separate group of chemolithoautotrophic bacteria

known as nitrite-oxidizing bacteria (NOB) (Ward 2008). Both oxidation steps require

molecular oxygen. Nitrification is thus an obligatory aerobic pathway.

Ammonia-oxidizing bacteria

Ammonia-oxidizing bacteria are ubiquitous in soils, freshwater, and marine

environments (Koops & Pommerening-Röser 2005). They are found exclusively in

three groups of Proteobacteria: the beta-proteobacterial Nitrosomonas and Nitrosospira,

and the gamma-proteobacterial Nitrosococcus (Head et al. 1993, Purkhold et al. 2000).

AOB metabolize NH4+ in the form of NH3 and oxidize it to NO2

− in a two-step process

(Figure 5, Kowalchuk & Stephen 2001). NH3 is first oxidized to hydroxylamine

(NH2OH) by the membrane-bound, multisubunit enzyme ammonia monooxygenase

(AMO). In the second step, NH2OH is oxidized to NO2− via the periplasmic enzyme

hydroxylamine oxidoreductase (HAO) (Arp et al. 2002). This second oxidation step

releases four electrons, of which two are returned to AMO and the other two are passed

23


via an electron transport chain to the terminal oxidase, thereby generating an

electrochemical H+ gradient over the cytoplasmic membrane (Figure 5). This proton

motive force is used for ATP-synthesis and for the formation of NADH by reverse

electron flow and provides the energy and reducing equivalents for CO2 fixation (Arp &

Stein 2003, Ferguson et al. 2007).

Periplasm

Cytoplasm

HAO

AMO

Periplasm

Cytoplasm

Periplasm

Cytoplasm

HAO

AMO

Figure 5: A scheme for electron transport pathways in the ammonia-oxidizing bacterium Nitrosomonas europaea. It is assumed that electrons derived from NH2OH are delivered from NH2OH dehydrogenase (HAO) to ubiquinone (UQ) via cytochrome c554 and cytochrome cm. It is assumed that cytochrome cm catalyses ubiquinol formation with concomitant uptake of H+ from the periplasmic side. The active site of the ammonia mono-oxygenase (AMO) is positioned on the periplasmic side. Protons released upon the putative oxidation of UQH2 by AMO are not shown but probably are released to the periplasm. Modified after Ferguson et al. (2007).

AOB are currently recognized as the main N2O producers under oxic conditions (Kool

et al. 2011). They produce N2O as a by-product during the oxidation of NH2OH to NO2−

(Figure 1 and 6). This is especially the case when the turnover of AMO and HAO are

not in balance and the concentration of NH2OH is increased (Cantera & Stein 2007, Yu

et al. 2010). The produced NH2OH can be reduced to NO and N2O either enzymatically

via HAO and NOR or chemically via chemodenitrification (Hooper & Terry 1979,

Stuven et al. 1992, Stein 2011).

24


Apart from this aerobic hydroxylamine oxidation pathway, AOB were shown to

produce N2O via a second distinct pathway, namely nitrifier denitrification. In this

process, AOB oxidize NH3 to NO2− that is subsequently reduced to NO, N2O and N2

(Ritchie & Nicholas 1972, Poth & Focht 1985, Bock et al. 1995). The reduction of NO2−

is analogous to that in canonical denitrification, with the denitrification enzymes NIR,

NOR and possibly also NOS being involved (Wrage 2001). Genes encoding the copper-

containing NIR (nirK) and the small and large subunits of NOR (norB and norC) have

been identified in various AOB strains, and it has been suggested that nitrifier

denitrification is a universal trait in the beta-proteobacterial AOB (Casciotti & Ward

2001, Schmidt et al. 2004, Shaw et al. 2006, Cantera & Stein 2007). However, genes

encoding the canonical NOS reductase of denitrifying bacteria have not been identified

in AOB genomes yet (Kim et al. 2010).

NH3

NH2OH

NO3�� NO2

��NAR NO N2O N2

NIR NOR NOS

NO2��

NO N2OHAO NOR

NO N2O N2NIR NOR NOS

HAO

AMO

NXR

NAP

Hydroxylamine oxidation

Nitrifier denitrification

Nitrite-oxidizing

bacteria (NOB)

Ammonia-oxidizing bacteria (AOB)

Denitrification

Denitrifying bacteria

NH3

NH2OH

NO3�� NO2

��NAR NO N2O N2

NIR NOR NOS

NO2��

NO N2OHAO NOR

NO N2O N2NIR NOR NOS

HAO

AMO

NXR

NAP

Hydroxylamine oxidation

Nitrifier denitrification

Nitrite-oxidizing

bacteria (NOB)

Ammonia-oxidizing bacteria (AOB)

Denitrification

Denitrifying bacteria

Figure 6: Major N2O producing pathways of nitrifying and denitrifying bacteria. Ammonia-oxidizing bacteria produce N2O via the hydroxylamine oxidation and the nitrifier denitrification pathway. Dashed lines indicate necessary reduction steps to consume N2O. AMO, ammonia monooxygenase; HAO, hydroxylamine oxidoreductase; NXR, nitrite oxidoreductase of nitrite-oxidizing bacteria (NOB); NAR and NAP, different types of nitrate reductase; NIR, nitrite reductase; NOR, nitric oxide reductase; NOS, nitrous oxide reductase. Modified after Stein (2011).

25


Although a rising number of studies suggests that nitrifier denitrification significantly

contributes to N2O production from soil, its verification remains difficult because of

methodological constraints (Kool et al. 2011 and references therein). AOB from pure

cultures and complex biofilms were shown to produce high amounts of N2O under low

O2 concentrations and/or high NO2− concentrations (Beaumont et al. 2004, Shaw et al.

2006, Schreiber et al. 2009). It is therefore suggested that the nitrifier denitrification

pathway is used to gain energy from the reduction of NO2− at O2-limiting conditions or

it is used to detoxify NO2− that is produced during nitrification (Poth & Focht 1985,

Bock et al. 1995, Beaumont et al. 2004).

Ammonia-oxidizing archaea

The oxidation of NH4+ to NO2

− was for more than a century exclusively attributed to

chemolithoautotrophic bacteria (Pester et al. 2011). The recent discovery of ammonia-

oxidizing archaea (AOA) within the novel phylum Thaumarchaeota radically changed

the view on the microbiology of nitrification (Konneke et al. 2005, Pester et al. 2011).

Metagenomic surveys targeting archaeal 16S rRNA genes and ammonia

monooxygenase genes (amoA) revealed the widespread distribution of archaea with the

potential capacity to oxidize NH3 as well as their numerical dominance over AOB in

many marine and terrestrial environments (Francis et al. 2005, Leininger et al. 2006,

Wuchter et al. 2006). These studies provide increasing evidence for the importance of

AOA in global biogeochemical cycles, but our knowledge about physiology and

ecosystem function of AOA is still in its infancy (Pester 2011). The biochemistry of

archaeal NH3 oxidation was proposed to be distinctively different from bacterial NH3

oxidation. Walker et al. (2010) suggested that AOA either use different enzymes for

NH3 oxidation via NH2OH to NO2−, or oxidize NH3 via nitroxyl (HNO) to NO2

−. The

latter hypothetical pathway would suggest that AOA do not produce N2O (Schleper &

Nicol 2010). However, Santoro et al. (2011) showed that AOA indeed produce N2O and

proposed that N2O production most likely arises from a process akin to nitrifier

denitrification. The authors conclude that AOA could play an important role in N2O

production in the near-surface ocean. Whether the ubiquitous AOA significantly

contribute to N2O production in terrestrial and marine environments awaits further

investigation.

26


Nitrite-oxidizing bacteria

Nitrite-oxidizing bacteria oxidize NO2− to NO3

− by the nitrite oxidoreductase (NXR)

(Figure 6). This oxidation step does not produce N2O. The resulting transfer of two

electrons to the terminal electron acceptor O2 generates a proton gradient that is used for

the synthesis of ATP and NADH reducing equivalents for CO2-fixation (Freitag & Bock

1990). The NO3− produced by NOB can be used directly by denitrifying bacteria in

environments with oxic-anoxic transition zones that allow nitrification and

denitrification to occur in close proximity (Figure 6).

Other pathways of nitrous oxide production

Besides N2O production through autotrophic nitrification, canonical denitrification and

nitrifier denitrification, several other N2O-producing processes exist. NO detoxification

pathways are broadly distributed throughout the bacteria and result in N2O production

via the enzymes flavohemoglobin (Hmp) and flavorubredoxin (NorVW) (Gardner et al.

2003, Stein 2011). Furthermore, ammonia-oxidation is not restricted to autotrophic

organisms, but can also be performed by a wide range of heterotrophic nitrifiers that do

not gain energy from the oxidation of NH3 (Robertson & Kuenen 1990, Wrage 2001).

Methane-oxidizing bacteria (MOB), which are closely related to AOB (Holmes et al.

1995), have also been shown to aerobically oxidize NH3 to NO2−, thereby releasing NO

and N2O (Campbell et al. 2011). DNRA is thought to produce low amounts of N2O as a

by-product (Kaspar & Tiedje 1981, Kelso et al. 1997, Cruz-Garcia et al. 2007) and is

increasingly recognized to be an important process in various environments (Silver et al.

2001, Lam et al. 2009, Koop-Jakobsen & Giblin 2010, Schmidt et al. 2011). In soils, for

instance, DNRA is suggested to significantly contribute to N2O production (Senga et al.

2006, Baggs 2011). So far, there is no evidence that N2O is produced by the anammox

process itself (Strous et al. 2006, Kuenen 2008, van der Star et al. 2008). However,

anammox bacteria were shown to produce trace amounts of N2O probably by

detoxification of NO, activity of NH2OH reductase or by reducing NO3− to NH4

+ (Kartal

et al. 2007). The potential significance of these diverse pathways as N2O sources in

various environments is, however, still unknown and needs further investigations (Stein

2011).

27


Environmental factors influencing nitrous oxide production

The amount of N2O released by nitrification and denitrification depends on the total rate

and the N2O yield of the process (fraction of N2O produced per NH4+ or NO3

−

consumed). Both are regulated by a suite of physiological and ecological factors, of

which oxygen concentration and substrate availability are of particular importance

(Stein 2011).

Low O2 concentrations generally lead to high N2O yields from both nitrification and

denitrification (Goreau et al. 1980, Betlach & Tiedje 1981, Codispoti 2010). The N2O

yield from nitrification increases with decreasing O2 concentration mainly because of an

increased rate of nitrifier denitrification (Wrage 2001 and references therein). The N2O

production from nitrification is usually favoured at hypoxic conditions when the N2O

yield is increased and enough O2 is present to sustain a reasonably high process rate

(Codispoti 2010). In denitrification, the increase in N2O yield under low O2 conditions

is due to decreased activity of the oxygen-sensitive N2O reductase (Bonin & Raymond

1990). The overall rate of denitrification is likely to slow down with increasing oxygen

concentration, as the facultative anaerobic denitrifiers will prefer O2 over NO3− as

terminal electron acceptor (Bonin & Raymond 1990). Therefore, the N2O production

from denitrification is particularly high under close to anoxic conditions when the

process rate of denitrification is still high and the N2O yield is increased compared to

completely anoxic conditions. Very high N2O production rates were observed for AOB

and denitrifers in pure cultures, a nitrifying reactor system, and an artificially grown

biofilm under rapidly changing oxygen conditions (Kester et al. 1997, Bergaust et al.

2008, Schreiber et al. 2009). The transient increase in N2O production was due to

unbalanced enzyme activity of AOB and denitrifiers in response to the shifts in oxygen

concentration.

Increased substrate availability generally stimulates the rates of nitrification and

denitrification according to Michaelis-Menten-kinetics (Barnard et al. 2005, Canfield et

al. 2005). Furthermore, high concentrations of NO3− can increase the N2O yield of

denitrification, if NO3− is preferred as an electron acceptor over N2O or even inhibits the

N2O-reductase (Blackmer & Bremner 1978, Firestone et al. 1979, Gaskell et al. 1981).

28


The rate and N2O yield of denitrification are also highly dependent on the availability of

electron donors. Increased availability of Corg stimulates the denitrification rates of

heterotrophic bacteria (Stehfest & Bouwman 2006). Furthermore, the balance between

the available electron donor and acceptor is important. A low C/NO3− ratio increases the

N2O yield, as Corg limits the final reduction step of denitrification (Knowles 1982,

Tiedje 1988, Morley & Baggs 2010).

Other important environmental factors that affect N2O production from nitrification and

denitrification are concentrations of intermediates, pH, and temperature (Stein & Yung

2003, Stein 2011). Elevated concentration of NO2−, for instance, increases N2O

production from AOB in microbial biofilms and from denitrification in an eutrophic

estuary (Dong et al. 2002, Schreiber et al. 2009). With decreasing pH, N2O production

increases from both nitrification and denitrification, as was, for instance, shown in soils

and riparian zones (van den Heuvel et al. 2011, Stehfest & Bouwman 2006, Richardson

et al. 2009). Temperature is generally a key factor controlling the metabolic rate of

microorganisms. The temperature response of nitrifiers and denitrifiers are bell-shaped

with typically highest metabolic rates between 20 and 35°C for mesophilic species

(Barnard et al. 2005 and references therein). In many environments, an increase in the

rates of nitrification, denitrification, and N2O emission has been observed with

increasing temperature (Avrahami et al. 2002, Braker et al. 2010).

Environmental controlling factors directly affect the process rates and N2O yields of

nitrification and denitrification through the short-term response of the existing microbial

communities. In addition, these factors also act as long-term environmental drivers and

influence the abundance and composition of microbial communities (Wallenstein et al.

2006). Since different nitrifier and denitrifier species can vary in their response to

environmental factors (e.g., different induction patterns of gene expression, enzyme

kinetics, and O2-dependence of N2O formation), the composition of nitrifying and

denitrifying communities have an impact on N2O production rates (Zumft 1997,

Cavigelli & Robertson 2000, Bange 2008). This impact is, however, difficult to resolve

and molecular investigations are needed to shed light upon the relationship between

microbial community structure and N2O production rates in natural environments.

29


Nitrogen cycling and nitrous oxide production in aquatic environments

Nitrogen cycling in aquatic environments

The nitrogen turnover rates in shallow aquatic environments are particularly high

compared to process rates in the open oceans or deep oceanic sediments. In shallow

aquatic systems, particulate organic nitrogen is built up by both pelagic and benthic

primary producers that assimilate NH4+ or NOx

−, or fix N2 (Figure 7). Benthic and

pelagic processes in these shallow aquatic environments are tightly coupled due to their

close proximity (MacIntyre et al. 1996). A large fraction of the PON from the water

column can thus reach the sediment surface (Suess 1980, Ferron et al. 2009). In addition,

freshwater and coastal ecosystems receive PON and especially DIN (mainly in the form

of nitrate) from terrestrial ecosystems by runoff (Boyer et al. 2006, Seitzinger et al.

2006, Schlesinger 2009). Due to intensive use of fertilizers on agricultural land, the

input of reactive nitrogen can be enormous, leading to very high nitrate concentrations

in the water column of many freshwater and coastal ecosystems (van Beusekom et al.

2008, Schlesinger 2009). Hence, sediments in freshwater, estuaries, and continental

shelf environments are characterized by high concentrations of organic and inorganic

nutrients, which sustain a dense and diverse community of organisms (Beukema 1991,

Herbert 1999). The benthic heterotrophic organisms play a key role in mineralizing

PON in the sediment and supplying inorganic nutrients for the benthic as well as pelagic

community (Herbert 1999, Nixon & Buckley 2002). Despite the high mineralization

rates, NH4+ rarely occurs at high concentrations in oxic environments, as it is either

readily re-assimilated into biomass, or oxidized to NO3− by nitrification (Canfield et al.

2005, Ward 2008).

Nitrification, a strictly aerobic process, only occurs in oxygenated water columns and in

oxic surface layers of sediments (Figure 7). The rates of nitrification reported for the

open ocean are in the range of a few to a few hundred nmol L−1 day−1, whereas the rates

in sediments, intertidal biofilms, and the water column of estuaries are often in the

range of μmol to mmol per m2 or L and day due to higher numbers of nitrifiers and

higher nutrient concentrations than in the oceans (Henriksen & Kemp 1988, de Wilde &

30


de Bie 2000, Magalhaes et al. 2005, Ward 2008). The depth to which nitrification

occurs in sediments is constrained by the limits of downward O2 diffusion, which is

typically a few mm depending upon sediment type, organic matter content, benthic

photosynthesis, and degree of mixing and bioturbation (Revsbech et al. 1980, Herbert

1999).

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Figure 7: Nitrogen cycling in aquatic environments showing the major N transformations and the N2O producing processes within the water column and the sediment. Solid lines indicate biological conversions of N compounds. Dashed lines indicate transport processes via diffusion or mixing by advection or bioturbation. Red lines represent N transformation and N2O emission by nitrification. Blue lines represent N transformation and N2O emission during denitrification. Nitrification prevails in the oxic water column and the upper oxic sediment layer, whereas denitrification prevails in oxygen-deficient water masses and deeper anoxic sediment layers. Other processes involved: (a) N2 fixation, (b) assimilation of NO3

− and NO2−, (c) NH4

+

assimilation, (d) remineralization, (e) DNRA, (f) anammox. DNRA and anammox can also prevail in hypoxic waters, but have so far not been identified as significant N2O source.

31


The dissimilatory reduction of nitrate can be fuelled by the supply of NO3− from

nitrification or from the water column and requires transport of NO3− into anoxic zones

or temporal separation of oxic production and anoxic consumption of NO3− (e.g., day-

time NOx− production followed by night-time dissimilation) (Seitzinger et al. 2006).

Denitrification, DNRA and anammox therefore prevail at high rates in environments

with oxic-anoxic interfaces (in space or time), such as aquatic sediments or hypoxic

zones in otherwise oxic water columns (Figure 7).

Denitrification is especially important in coastal areas, where it removes a large fraction

of terrestrial DIN inputs as N2 gas (Seitzinger & Kroeze 1998, Galloway et al. 2004,

Seitzinger et al. 2006). The rates of denitrification in the sediment typically range from

0.1 to 10 mmol m−2 d−1 (Joye & Anderson 2008). In sediments, the coupling of

nitrification and denitrification can be very close and nitrification can supply up to

100% of the NO3− consumed by denitrification (Ward 2008). The water-column NO3

− is

especially an important driver for sedimentary denitrification in eutrophic environments

where high NO3− concentrations in the water column support NO3

− diffusion into the

sediment (Joye & Anderson 2008). In oceanic oxygen minimum zones, denitrification

rates are in the range of nanomolar per day (Lam & Kuypers 2011).

Nitrous oxide production in aquatic environments

Nitrification and denitrification are recognized as the primary N2O-producing processes

in aquatic environments (Ivens et al. 2011). Their relative importance for aquatic N2O

emissions is, however, still a matter of debate. This is mainly due to the facts that

experimental studies on aquatic emissions of N2O are scarce and quantification is

complicated due to the complex network of N cycling processes, the close proximity of

nitrification and denitrification activities in sediments, and the large spatial and

temporal variability of N2O emissions (Gruber 2008, Ivens et al. 2011). Sedimentary

denitrification and water-column nitrification seem to be the major N2O-producing

processes in coastal areas (Bange 2006, 2008), while in the open oceans, the majority of

N2O emissions is attributed to water-column nitrification (Suntharalingam & Sarmiento

2000, Nevison et al. 2003, Bange 2008).

32


The N2O yields from nitrification and denitrification in aquatic systems are usually

lower than 1% (Seitzinger 1988, Bange 2008). High N2O production rates therefore

occur only in environments with high process rates of nitrification and denitrification.

How much of the N2O produced in aquatic environments is finally emitted to the

atmosphere depends on the distance of the N2O production site to the atmosphere and

the prevailing hydrodynamics. Consequently, freshwater and coastal areas as sites of

high N turnover in close proximity to the atmosphere have much higher areal N2O

emission rates than the open oceans (Seitzinger et al. 2000, 2006). Therefore, they

significantly contribute to the global aquatic N2O emission despite their relatively small

surface area. Intense sites of N2O production in the oceans are oxygen minimum zones

(Codispoti et al. 2001). They make up less than 1 % of the ocean’s volume, but they are

estimated to account for 25−50% of the total oceanic N2O emissions (Suntharalingam &

Sarmiento 2000). Highest rates of N2O emission occur in coastal upwelling regions and

estuaries where N2O production is stimulated due to very high nutrient concentrations

and oxygen-deficient conditions close to the water surface (Codispoti 2010, Naqvi et al.

2010). Here, N2O supersaturations of up to 8000% at the water-atmosphere interface

result from the upwelling of subsurface water (Naqvi 2000, Bange 2006, Naqvi et al.

2010). It is assumed that periodic aeration due to turbulence in these shallow hypoxic to

anoxic zones leads to “stop and go” denitrification. The frequent changes in oxygen

concentration result in the accumulation of N2O due to a more pronounced inhibition of

the N2O reductase by oxygen and/or due to a delayed expression of the N2O reductase

during the onset of denitrification (Naqvi et al. 2000, Codispoti et al. 2001).

The total volume of oxygen-deficient zones is expected to increase in the future due to

increased eutrophication leading to higher productivity and consequently higher O2

consumption during organic matter degradation (Diaz & Rosenberg 2008).

Anthropogenic nutrient inputs thus indirectly increase N2O emission from aquatic

environments by stimulating the rate of nitrification and denitrification and by causing

hypoxia in eutrophic regions and thereby extending the area of high N2O production by

denitrification.

33


Effects of benthic macrofauna on nitrogen cycling

Benthic macrofauna directly affect nitrogen cycling by ingesting PON and excreting

ammonium and feces (Christensen et al. 2000, Michaud 2006). Moreover, they also

indirectly influence biogeochemical nitrogen cycling by affecting the distribution,

metabolism and composition of microbial communities in aquatic environments (Harris

1993, Papaspyrou et al. 2006, Bertics & Ziebis 2009). The impact of macrofauna on

microbial nitrogen cycling is especially strong in aquatic sediments that are densely

inhabited by diverse communities of epi- and infaunal invertebrates (Cadée 2001).

These macrofaunal communities can significantly alter the physicochemical properties

of the sediment through their bioturbation, bioirrigation, and feeding activities, and

create a three-dimensional temporally and spatially dynamic mosaic of micro-

environments (Aller & Aller 1998, Kristensen 2000).

Bioturbation and bioirrigation

Invertebrates living inside the sediment (infauna) redistribute large amounts of sediment

and construct burrows and tubes deep into the sediment (bioturbation, Aller & Aller

1998). These structures enlarge the area of the sediment-water interface and extend the

oxic-anoxic transition zone into otherwise anoxic sediment layers (Reise 2002). They

thus increase the area for diffusive solute exchange between anoxic porewater and the

overlying water, and are sites of intense microbial colonization and activity (Papaspyrou

et al. 2006). Burrow-dwelling species further stimulate microbial metabolism by

periodically flushing and ventilating their burrows with the overlying water

(bioirrigation, Aller et al. 2001). Thereby, they introduce oxygenated water into deeper

sediment layers and enhance particle and solute fluxes across the sediment-water

interface. The resulting increased availability of oxygen, alternative electron acceptors,

inorganic nutrients, and organic matter in deeper sediment layers supports aerobic

metabolism, coupled redox reactions, and remineralization of fresh and aged PON

(Henriksen et al. 1983, Mortimer et al. 1999, Kristensen & Mikkelsen 2003, Wenzhofer

& Glud 2004).

34


The increased supply of O2 due to burrow ventilation and of NH4+ due to the animal’s

excretion and stimulated remineralization rates enhances nitrification in the burrow

environment. Increased nitrification in turn enhances dissimilatory nitrate reduction in

adjacent anoxic zones by the increased supply of nitrate (Pelegri & Blackburn 1994,

Nielsen et al. 2004, Stief & de Beer 2006). The lining and walls of burrows are thus

particularly favourable sites for coupled nitrification-denitrification. Furthermore, the

burrow walls are usually enriched in organic matter originating from mucus secretion

by the inhabitants and trapped detritus particles, and are characterized by fluctuating

redox conditions due to the periodic irrigation of the burrows (Nielsen et al. 2004,

Papaspyrou et al. 2006). Burrow walls therefore provide unique microenvironments that

often promote higher abundance and activity of N-converting microorganisms than the

oxic-anoxic interface in the ambient sediment.

The impact of benthic invertebrates on microbial activities and biogeochemical

processes varies with species and depends on the animal’s abundance, size, metabolic

activity, feeding type, mode of sediment mixing, irrigation, and biogenic structure

building (Mermillod-Blondin & Rosenberg 2006). Species that modify the physical

environment and thereby regulate the availability of resources for other organisms are

referred to as ecosystem engineers (Jones et al. 1994). One of the most important

ecosystem engineers in temperate coastal marine sediments is the common ragworm

Hediste diversicolor, formerly known as Nereis diversicolor (Figure 8, Kristensen

2001). This abundant polychaete strongly alters the physicochemical properties of

intertidal sediments by its vigorous burrow ventilation, contributes to mechanical

breakdown and mixing of PON as deposit feeder, and increases the input of organic

matter to the sediment surface when filter feeding. These worms consequently affect the

entire biological community in soft-bottom habitats and play an important role in

enhancing organic matter decomposition and removal of bioavailable nitrogen through

simultaneous stimulation of nitrification and denitrification (Kristensen & Mikkelsen

2003, Nielsen et al. 2004, Papaspyrou et al. 2006). In Chapter 6 of this thesis, it is

shown that this polychaete also exerts a strong control on the pool of intracellular nitrate

in intertidal sediments, and thus on the fate and availability of nitrogen in benthic

systems.

35


a ba b

Figure 8: (a) The polychaete Hediste diversicolor, (b) cross section of sediment bioturbated by H. diversicolor. Light patches are oxidized sediment surrounding older burrows. Note that the worm in the centre inhabits a newly constructed burrow without noticeable oxidized sediment. Panel b was taken from Kristensen (2001).

Colonization of benthic invertebrates

The internal and external surfaces of benthic invertebrates can be colonized by

microorganisms (Wahl 1989, Carman & Dobbs 1997, Welsh & Castadelli 2004).

Especially the hard external surface of invertebrates that live on the sediment surface or

on hard substrates (epifauna) can be covered by thick microbial biofilms. Among the

dominant epifaunal species in coastal regions that provide such colonization surfaces is

the blue mussel Mytilus edulis (Figure 9, Bouma et al. 2009). This common bivalve

forms stable, permanent beds with individuals attached to each other by byssus threads.

Its shell surface serves as habitat for a highly diverse epibiont community, including

many microorganisms (Asmus 1987, Dittmann 1990). The mussel beds are highly

enriched in nutrients due to the efficient filter-feeding and high ammonium excretion

rate of the mussel (Prins et al. 1996, Smaal & Zurburg 1997).

Like M. edulis, many invertebrate species excrete high amounts of ammonium and

thereby significantly contribute to the overall NH4+ production in sediments (Blackburn

& Henriksen 1983, Dame & Dankers 1988, Smaal & Zurburg 1997). By supplying

ammonium and providing a colonization surface, invertebrates represent a suitable

habitat for nitrifying bacteria (Welsh & Castadelli 2004). Indeed, high potential

nitrification rates were found for different epi- and infaunal invertebrate species (Welsh

& Castadelli 2004). Increased nitrogen turnover seems thus not only to be linked to

36


microbial activity in the animal’s burrows, but also to microbial activity on the surfaces

of invertebrates. However, studies on nitrogen cycling in biofilms on external surfaces

of invertebrates are scarce. This thesis therefore investigated for the first time the

potential and mechanisms of microbial N2O production in exoskeletal biofilms on

aquatic invertebrates (Chapters 2 to 4).

Figure 9: The Blue Mussel Mytilus edulis, (a) small part of the mussel colony in which individuals are attached to each other by byssus threads, (b) the shell surface of M. edulis is usually colonized by a diverse community of epibionts, (c) close-up on the biofilm community, (d) confocal laser scanning microscope picture of the microbial community in the shell biofilm of M. edulis, overlay of the top 400 μm of the biofilm (blue: DAPI staining, green: chlorophyll, red: phycocyanin).

Trophic interactions

Macrofauna also directly affect the biomass, activity and community composition of

microbes by ingesting free-living or particle-attached microorganisms (Plante & Wilde

2004). Many invertebrates possess enzymes in their gut to lyse and digest at least part of

the ingested bacteria and use them as a food source (McHenery & Birkbeck 1985,

Plante & Shriver 1998). The ingestion of microbes by invertebrates is, however, often

not a simple consumption of food. Different functional and phylogenetic groups of

37


microorganisms experience different fates during gut passage (Harris 1993). Depending

on their ability to resist digestion and adapt to the prevailing conditions in the animal’s

gut, subgroups of ingested microbes might be lysed, survive, get metabolically activated

or even grow during gut passage (Plante & Jumars 1992, Harris 1993). The gut

microenvironment of invertebrates can be very distinct from the ambient environment

from which the microbes were ingested. The gut of invertebrates can be anoxic or O2-

limited and enriched in nutrients (Horn et al 2003, Plante & Jumars 1992, Stief & Eller

2006). These guts provide suitable habitats for anaerobic microbes such as denitrifying

bacteria that will remain or become metabolically active while passing the gut or even

permanently colonize the gut (Drake & Horn 2007). The viable microbes can be of

advantage for the host by producing exoenzymes that help digesting complex organic

matter (Harris 1993). On the other hand, viable microbes in the gut can also be of

disadvantage if they compete with the host for limiting nutrients (Harris 1993, Drake &

Horn 2007). As a consequence of these different responses of ingested microbes to the

specific conditions in the invertebrate gut, the gut passage leads to changes in the

composition and activity of the microbial community compared to that found in the

surrounding sediment (Harris 1993).

Nitrous oxide emission from macrofauna

Biogenic N2O emission is classically linked to microbial activities in soils, sediments

and water bodies. However, in addition to these sources, N2O emission was also

reported for earthworms and freshwater invertebrates (Karsten & Drake 1997, Stief et al.

2009). These animals host microorganisms that produce N2O and are thus N2O-emitters,

but not N2O-producers.

Nitrous oxide emission from earthworms

Different earthworm species were found to emit N2O with an average rate of 1.5 nmol

g−1 (fresh weight) h−1, and global N2O emission from earthworms were estimated to be

0.19 Tg N yr−1 compared to the total global emission of 17.7 Tg N yr−1 (Drake et al.

2006, Drake & Horn 2007, Denman et al. 2007). The emission rates on a dry weight

38


basis can be far higher from earthworms than from bulk soils (Depkat-Jakob et al. 2010

and references therein) and earthworms can contribute up to 56% of the in situ N2O

emission from certain soils (Karsten & Drake 1997, Matthies et al. 1999, Borken et al.

2000). The N2O emission by earthworms was found to be due to the activation of

ingested denitrifiers by the specific in situ conditions in the gut (Drake & Horn 2006,

Horn et al. 2006). In contrast to the ambient soil, the gut microenvironment is

characterized by anoxia, high water content, high concentration of readily degradable

Corg, and presence of NO3− and NO2

− (Figure 10, Horn et al. 2003, Drake et al. 2006).

The earthworm gut thus constitutes a unique microsite in aerated soils that provides

ideal conditions for denitrifiers and other microorganisms capable of anaerobic growth

(Drake & Horn 2007). Denitrification in the gut of earthworms results in the emission of

about equal amounts of N2O and N2 (Drake & Horn 2007). It has been suggested that

this very high N2O yield is due to a delay in the synthesis of N2O reductase or high

concentrations of NO2− (Horn et al. 2003, Ihssen et al. 2003). Although soil-derived

denitrifiers are recognized as the main N2O-producing microbes in the earthworm gut,

non-denitrifying dissimilatory NO3- reducers might indirectly contribute to N2O

production by providing high concentrations of NO2-, which has been shown to

stimulate N2O production more effectively than NO3- (Matthies et al. 1999, Ihssen et al.

2003, Drake & Horn 2007).

Figure 10: Hypothetical model illustrating which factors stimulate the production of N2O and N2 by bacteria ingested into the earthworm gut. The relative concentrations of compounds are reflected in the font size, and the relative effect of each compound on the production of N2O and N2 in the gut is indicated by the thickness of the arrows. The main factors that appear to stimulate ingested denitrifiers in the gut are in red. Taken from Drake & Horn (2007) who modified the scheme after Horn et al. (2003) and Drake et al. (2006).

39


Nitrous oxide emission from freshwater invertebrates

Similar to earthworms, diverse freshwater invertebrate species were found to emit N2O

with emission rates ranging from 0 to 93.1 nmol g−1 (dry weight) h−1 (Stief et al. 2009).

N2O emission was, like for earthworms, ascribed to the denitrification activity of

ingested bacteria in the anoxic animal gut. The N2O emission rates of freshwater

invertebrates largely depend on the amount of ingested bacteria, which is influenced by

the animal’s diet. Filter feeders and deposit feeders that prefer a bacteria-rich detritus

diet show the highest rates, shredders and grazers intermediate, and predators ingesting

a bacteria-poor carnivorous diet very low N2O emission rates (Stief et al. 2009, Stief &

Schramm 2010). Like in earthworms, the N2O yield of denitrification in the guts of two

abundant filter- and deposit-feeding insect larvae were exceptionally high, ranging from

15 to 68% of the N gas flux (Stief et al. 2009). Since aquatic filter- and deposit-feeders

typically ingest particle-attached or free-living bacteria from the oxic water column or

oxic surface sediment layer, bacteria probably experience an oxic-anoxic shift when

being ingested into the anoxic animal gut. It is hypothesized that this shift activates

ingested facultative denitrifiers and that during the onset of denitrification the induction

of the N2O reductase is delayed, leading to the accumulation and emission of N2O from

the animal gut (Figure 11, Stief et al. 2009).

In addition to direct stimulation of N2O production in the animal gut, burrowing

invertebrates were shown to enhance N2O and N2 emission from the surrounding

sediments (Figure 11, Svensson 1998, Stief et al. 2009, Stief & Schramm 2010). This

indirect stimulation of N2O emission is probably due to the animal’s bioirrigation

activity causing periodic changes between oxic and anoxic conditions in the burrows

and increased nutrient supply, thus enhancing the rates of N transformation and N2O

production. The excretion of fecal pellets that contain active denitrifiers might further

enhance the capacity of the sediment to produce N2O. Invertebrates that do not emit

N2O themselves may thus indirectly contribute to the stimulation of N2O emission from

sediments by their bioirrigation activities (Stief & Schramm 2010). On the other hand,

N2O emitted by infaunal species might be partially consumed by denitrification in the

surrounding sediment. For infaunal invertebrates, the sediment might thus acts as an

additional source or sink of N2O (Stief et al. 2009, Stief & Schramm 2010).

40


Figure 11: Conceptual model of the activation of ingested denitrifying bacteria in the gut of aquatic invertebrates and the resulting enhancement of N2O emission from sediment. (1) Bacteria in the water column or in the surface sediment are exposed to oxic conditions and therefore do not exhibit NO3

− reduction activity (blue ovals). (2) Invertebrates that feed on organic particles to which bacteria are attached transfer the bacteria to anoxic conditions in their guts. Anoxia and the presence of NO3

− lead to the activation of ingested denitrifying bacteria in the gut of the invertebrate (orange ovals) that reduce NO3

− to N2O and N2. The produced N2O diffuses through the gut wall of the invertebrates into the surrounding sediment or is pumped out of the tube by ventilation activity of the invertebrate. (3) Animal burrows in which O2 and NO3

− concentrations fluctuate (stippled circles) are inoculated with actively NO3−-reducing

bacteria. As a consequence, NO3− reduction and concomitant N2O production in animal guts and

in animal-influenced sediment are higher than in non-inhabited sediment. Kindly provided by P. Stief.

Important environmental factors that influence N2O emission from freshwater

invertebrates are the ambient NO3− concentration and the temperature (Stief et al. 2010,

Stief & Schramm 2010). However, one of the factors must exceed a certain threshold

value before the other factor can stimulate N2O emission. For instance, N2O emission

rate of the insect larvae Chironomus plumosus is only increased by temperature when

the NO3− concentration exceeds 25−50 μmol L−1, and by NO3

− when the temperature is

above 4−10°C (Stief et al 2010). Accordingly, rates of N2O emission from this insect

larvae vary seasonally depending on the prevailing temperature and NO3− concentration

like it is known for denitrification rates in sediments (Jørgensen & Sorensen 1985,

Jørgensen & Sorensen 1988). In temperate freshwater and coastal waters, NO3−

41


availability and temperature are mostly antagonistic during the year with highest NO3−

concentrations in winter and lowest concentrations in summer. Highest rates of

sedimentary denitrification and N2O emission are therefore observed in spring and

autumn when moderate NO3− concentrations coincide with moderate temperatures. N2O

emission rates of freshwater invertebrates appear to undergo the same seasonal changes

as typically found in aquatic sediments unless other factors (e.g., larval development or

type and rate of feeding) limit N2O production in the gut of the animal (Stief &

Schramm 2010).

Nitrous oxide emission from other organisms

Besides benthic macrofauna, a variety of other organisms were shown to emit N2O.

High amounts of N2O are produced by cattle production. However, the majority of this

N2O derives from microbial processes in the animal waste (Oenema et al. 2005). The

bovine digestive track may be only a very small source of N2O, since here dissimilatory

nitrate reduction to ammonium takes place which might produce trace amounts of N2O

that could escape to the atmosphere during rumination (Kaspar & Tiedje 1981). In a

contributed work, it is shown that N2O is produced in the human oral cavity via

denitrifying bacteria in the dental plaque. The average rate of oral N2O emission was 80

nmol h−1 per individual. Extrapolated to the world population, humans produce about

0.00013 Tg N yr−1, which represents a rather insignificant amount of the global annual

N2O emissions. In contrast, the N2O emission by plants might be of global importance

despite their relatively low N2O emission rates because of their huge biomass (Smart &

Bloom 2001). The ability to produce N2O seems to be widespread among different plant

species (Smart & Bloom 2001, Hakata et al. 2003). Very recently, also two different

soil-feeding termite species were shown to emit N2O (Ngugi & Brune 2012). The N2O

and N2 emission rates per gram fresh weight are in the same range as those reported for

earthworms, and their production was associated with the nutrient-rich gut of the

animals. Denitrification mainly takes place in the posterior hindgut, while dissimilatory

reduction of NO3− to NH4

+ occurs throughout the gut at far higher rates than

denitrification. It is hypothesized that both denitrification and DNRA might be involved

in N2O emission from termites, since N2O is not only produced in the posterior hindgut,

but also in other sections of the termite gut (Ngugi & Brune 2012).

42


The list of microbial processes that produce N2O and of organisms that act as N2O

emitters is long. However, marine invertebrates, which densely colonize marine

sediments, have not been investigated so far.

43

Chapter 1 Aims of the thesis

Aims of the thesis

This thesis aimed at improving our knowledge about invertebrate-microbe interactions

and their role in biogeochemical nitrogen cycling and the production of the greenhouse

gas N2O in aquatic environments.

As outlined in the previous sections, macrofauna strongly interact with microbes and

thereby influence biogeochemical nitrogen cycling, leading to increased rates of

nitrification and denitrification, and concomitant production and emission of N2O to the

atmosphere. Terrestrial and freshwater invertebrates were reported to be emitters of this

important greenhouse gas. N2O emission by marine invertebrates, however, has not

been studied so far. The major focus of this thesis was therefore to unravel the potential

and the underlying mechanisms of microbial N2O production associated with marine

invertebrates. According to previous studies on terrestrial and freshwater macrofauna,

coastal benthic invertebrates were assumed to be suitable candidates for high N2O

emission potentials, since they inhabit nitrate- and organic-carbon-rich environments

and comprise many filter- and deposit-feeding species. Therefore, in two studies of this

thesis, macrofauna was collected from two coastal field sites that are representative for

many tidal and subtidal habitats in temperate regions: an intertidal flat in the German

Wadden Sea and the shallow Aarhus Bay in Denmark. Further studies included in this

thesis investigated the potential and mechanisms of N2O emission from a highly

abundant freshwater species, the Zebra Mussel Dreissena polymorpha, and an important

aquaculture species, the Pacific White Shrimp Litopenaeus vannamei, which is typically

reared under highly nutrient-enriched conditions and therefore was assumed to be a

strong N2O-emitter.

The main research questions of this thesis were:

� Do marine invertebrates emit N2O?

� Is the ability to emit N2O restricted to a certain taxonomic group or is it a

widespread trait among marine invertebrates?

� Which environmental factors and species characteristics influence the rates of

N2O emission?

44

Chapter 1 Aims of the thesis

� Does the N2O emission potential depend on the feeding guild like in freshwater

invertebrates?

� At which site(s) is the N2O produced in the animal body? Is it only produced in

the gut or are there other N2O-producing animal compartments? If yes, how

important are they?

� Which microbial N-cycling processes are involved in N2O production and how

much do they contribute to the total N2O emission?

� How does the gut microenvironment affect the composition and activity of

ingested microorganisms?

These research questions were addressed by a combination of (i) incubation and stable

isotope experiments with subsequent measurements of N2O emission rates by gas

chromatography or mass spectrometry, (ii) investigations of the animal-associated

microbial community by molecular analysis, and (iii) analysis of the driving

environmental factors by microsensor measurements and statistical analysis.

An additional study investigated the origin and ecological controls of a large

sedimentary pool of intracellular nitrate that was discovered at the sampling site in the

Wadden Sea. Since the sediment was densely colonized by the polychaete Hediste

diversicolor, it was hypothesized that the worm stimulates the formation of intracellular

nitrate by interacting with nitrifying bacteria and nitrate-storing diatoms through its

bioturbation activities. This hypothesis was investigated in a long-term microcosm

experiment.

45

Chapter 1 Overview of manuscripts

Overview of manuscripts

Chapter 2:

Nitrous oxide production associated with coastal marine invertebrates Ines M. Heisterkamp, Andreas Schramm, Dirk de Beer, Peter Stief

The study was initiated by P. Stief and A. Schramm. The concept and design of the

study was developed by P. Stief, I.M. Heisterkamp, and A. Schramm. I.M. Heisterkamp

performed sampling, rate measurements, and statistical analysis with help of P. Stief.

The manuscript was written by I.M. Heisterkamp with editorial help and input from the

co-authors.

The manuscript is published as Feature Article in Marine Ecology Progress Series 415:

1-9, 2010

Chapter 3:


processes, precursors and relative importance Ines M. Heisterkamp, Andreas Schramm, Lone H. Larsen, Nanna B. Svenningsen,

Gaute Lavik, Dirk de Beer, Peter Stief

The concept and experimental design of the study were developed by I.M. Heisterkamp,

P. Stief, and A. Schramm. The design of the stable isotope experiments was conceived

by P. Stief together with G. Lavik. I.M. Heisterkamp carried out the sampling and the

laboratory work including short-term and long-term incubation experiments, stable

isotope experiments, rate measurements with gas chromatography and mass

spectrometry, microsensor measurements, analysis of nutrients, protein contents and

ammonium excretion. L.H. Larsen and N.B. Svenningsen contributed to measurements

of the N2O emission rates and ammonium excretion rates and P. Stief to the stable

isotope experiments. Analysis and evaluation of the data was done by I.M. Heisterkamp.

G. Lavik helped with evaluating the mass spectrometry data. I.M. Heisterkamp

46


conceived and wrote the manuscript with input from P. Stief, A. Schramm, G. Lavik,

and D. de Beer.

The manuscript is accepted in Environmental Microbiology

Chapter 4:

Shell biofilm nitrification and gut denitrification contribute to emission of

nitrous oxide by the invasive freshwater mussel Dreissena polymorpha

(Zebra Mussel) Nanna B. Svenningsen, Ines M. Heisterkamp, Maria Sigby-Clausen, Lone H. Larsen,

Lars Peter Nielsen, Peter Stief, Andreas Schramm

I.M. Heisterkamp contributed to the development of the concept and experimental

design and helped with rate measurements.

The manuscript is published in Applied and Environmental Microbiology 78(12):4505-

4509, 2012

Chapter 5:

Incomplete denitrification in the gut of the aquacultured shrimp Litopenaeus

vannamei as source of nitrous oxide Ines M. Heisterkamp, Andreas Schramm, Dirk de Beer, Peter Stief

The study was conceived by I.M. Heisterkamp, P. Stief, and A. Schramm. All

experiments, rate measurements, and microsensor measurements were conducted by I.M.

Heisterkamp. The preliminary manuscript was written by I.M. Heisterkamp with

editorial help of P. Stief.

Chapter 5 presents the preliminary data on microbial N2O production associated with

the aquacultured shrimp Litopenaeus vannamei. Molecular analysis of the abundance

47


and expression of denitrification genes in different sections of the gut and in the water

of the aquaculture farm is still in progress.

Chapter 6:

Indirect control of the intracellular nitrate pool of intertidal sediment by the

polychaete Hediste diversicolorInes M. Heisterkamp, Anja Kamp, Angela T. Schramm, Dirk de Beer, Peter Stief

The concept and experimental design of the study were developed by P. Stief, I.M.

Heisterkamp, and A. Kamp. I.M. Heisterkamp was responsible for sampling of animals

and sediment, the set-up and implementation of the experiment, and nutrient analysis. A.

Kamp analyzed the intracellular nitrate and A.T. Schramm the photopigments.

Microsensor measurements, depth-integration of data and statistical analysis was done

by P. Stief. I.M. Heisterkamp wrote the manuscript mainly in collaboration with P. Stief

and input from A. Kamp and D. de Beer.

This manuscript is published in Marine Ecology Progress Series 445:181-192, 2012

48

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62

Chapter 2

Feature Article photograph: N2O production associated with the snail Hinia reticulata partly results from microbial activity in exoskeletal biofilms covering the shell.

63

Nitrous oxide production associated with coastal marine invertebrates

Ines Maria Heisterkamp1, Andreas Schramm2, Dirk de Beer1, and Peter Stief1

1Max Planck Institute for Marine Microbiology, Microsensor Group,

Celsiusstraße 1, D-28359 Bremen, Germany

2Department of Biological Sciences, Microbiology, Aarhus University,

Ny Munkegade 114, DK-8000 Aarhus C, Denmark

Feature Article in Marine Ecology Progress Series 415: 1-9, 2010

65

Chapter 2 N2O emission by marine invertebrates

Abstract

Several freshwater and terrestrial invertebrate species emit the greenhouse gas nitrous

oxide (N2O). The N2O production associated with these animals was ascribed to

incomplete denitrification by ingested sediment or soil bacteria. The present study

shows that many marine invertebrates also emit N2O at substantial rates. A total of 19

invertebrate species collected in the German Wadden Sea and in Aarhus Bay, Denmark,

and 1 aquacultured shrimp species were tested for N2O emission. Potential N2O

emission rates ranged from 0 to 1.354 nmol individual�1 h�1, with an average rate of

0.320 nmol individual�1 h�1, excluding the aquacultured shrimp Litopenaeus vannamei,

which showed the highest rate of N2O emission measured so far for any marine species

(3.569 nmol individual�1 h�1), probably due to very high nitrate concentrations in the

rearing tanks. N2O emitted by L. vannamei was almost exclusively produced in its gut

by incomplete denitrification. Statistical analysis revealed that body weight, habitat, and

exoskeletal biofilms were important determinants of animal-associated N2O production.

The snail Hinia reticulata emitted about 3.5 times more N2O with an intact exoskeletal

biofilm on its shell than with an experimentally cleaned shell. Thus, N2O production

associated with marine invertebrates is apparently not in every species due to gut

denitrification, but may also result from microbial activity on external surfaces of the

animals. The high abundance and potential N2O emission rates of many marine

invertebrate species suggest significant contributions to overall N2O emissions from

coastal marine environments and aquaculture facilities.

66


Introduction

Nitrous oxide (N2O) is the third most important greenhouse gas after carbon dioxide and

methane. Its atmospheric concentration is rapidly increasing, and it contributes

significantly to global warming (IPCC 2007) and to the depletion of the stratospheric

ozone layer (Ravishankara et al. 2009). Biogenic N2O emission originates primarily

from soils and oceans, where microbial nitrification and denitrification are the major

N2O-producing processes (Mosier et al. 1998, Stein & Yung 2003). During nitrification

(the 2-stage oxidation of ammonium to nitrate) N2O is produced as a byproduct in the

first oxidation step (Goreau et al. 1980), whereas in denitrification (the respiratory

reduction of nitrate or nitrite to nitrogenous gases) N2O is produced as a true

intermediate (Zumft 1997). The complete denitrification pathway involves 4 enzymes

that reduce nitrate to dinitrogen stepwise via the intermediates nitrite, nitric oxide, and

N2O. The 4 reductases are induced sequentially under anoxic conditions when oxidized

inorganic nitrogen compounds and appropriate electron donors are available (Tiedje

1988, Zumft 1997). Whether denitrification acts as a source or sink of N2O depends on

the presence and activity of nitrous oxide reductase, which shows a higher sensitivity

towards oxygen, lower carbon-to-nitrate ratios, and lower pH than the other 3 enzymes

(Tiedje 1988, Bonin & Raymond 1990).

Important sites of N2O emission are environments that are characterized by high input

and turnover rates of inorganic nitrogen, such as fertilized soils and coastal areas

(Mosier et al. 1998, Seitzinger & Kroeze 1998, Bange 2006). Microbial nitrogen

conversions and concomitant N2O production are especially stimulated in coastal

sediments and in rock biofilms, due to high riverine input of nitrogen (Seitzinger &

Nixon 1985, Law et al. 1992, Middelburg et al. 1995, Robinson et al. 1998, Magalhaes

et al. 2005). Nitrification activity prevails at the oxic sediment surface and is fuelled by

ammonium from organic matter degradation. Denitrification activity prevails in the

anoxic subsurface layer and is driven by nitrate from nitrification (i.e. coupled

nitrification–denitrification) or the water column (Jenkins & Kemp 1984). Sedimentary

denitrification is commonly assumed to be the major source of N2O to the water

column, with benthic N2O fluxes making up approximately 1% of the dinitrogen fluxes

(Seitzinger 1988, Magalhaes et al. 2007, Ferrón et al. 2009). Sedimentary nitrification

67


can, despite lower N2O production rates, significantly contribute to benthic N2O fluxes,

due to its proximity to the sediment surface (Meyer et al. 2008). Oversaturation of N2O

in the water column occurs in many coastal areas (Kieskamp et al. 1991, Middelburg et

al. 1995, Robinson et al. 1998, Dong et al. 2002).

Besides the microbial N2O production in soils, sediments, and water bodies, N2O is also

emitted by earthworms and freshwater invertebrates (Karsten & Drake 1997, Drake &

Horn 2007, Stief et al. 2009). This animal-associated N2O production is due to the

denitrification activity of ingested bacteria in the anoxic gut. The specific in situ

conditions of the earthworm gut, including anoxia and high concentrations of easily

degradable organic carbon, as well as nitrate or nitrite, stimulate the activity of ingested

N2O-producing soil bacteria (Drake et al. 2006). A similar mechanism has been

suggested for freshwater invertebrates, whose N2O emission is largely explained by

their preferred diet: filter- and deposit-feeders show high, shredders and grazers

intermediate, and predators very low N2O emission rates (Stief et al. 2009). This

suggests that N2O emission is caused by bacteria that are coingested with the food taken

up by freshwater invertebrates. N2O emission rates of both terrestrial and freshwater

invertebrates increase with nitrate and temperature and decrease with oxygen

availability, indicating the important role of these environmental factors for gut

denitrification (Karsten & Drake 1997, Matthies et al. 1999, Stief et al. 2009, 2010,

Stief & Schramm 2010).

The N2O emission potential of marine invertebrates has so far been neglected, although

coastal marine sediments are presumably hot spots of N2O emission, since they are

densely inhabited by filter- and depositfeeding invertebrates (Williams et al. 2004,

Philippart et al. 2007) and exposed to high nitrate concentrations (Kieskamp et al. 1991,

Van Beusekom et al. 2008). High N2O emission can also be expected from aquaculture

facilities in which animals are typically reared at high densities and high nitrate

concentrations. The present study, therefore, investigated the N2O emission potential of

different marine invertebrate species from coastal sediments of the North Sea and Baltic

Sea and of the aquacultured shrimp Litopenaeus vannamei. To understand how the N2O

emission potential of marine invertebrates is controlled by abiotic and biotic factors,

correlations between potential N2O emission rates and species-specific traits were

investigated by statistical analysis.

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Materials and Methods

Sampling of animals

We tested the N2O emission potential of 19 benthic invertebrate species from the

German Wadden Sea and the Aarhus Bay in Denmark, and of the aquacultured shrimp

Litopenaeus vannamei (provided by Ecomaris Marifarm Kiel, Germany). Sampling was

carried out between March and June 2008 at the mixed sediment intertidal flat near

Dorum-Neufeld (53°45'N, 8°21'E) and at 3 different sites in Aarhus Bay (56°9.75’N,

10°16.80’E; 56°9.29’N, 10°19.15’E; 56°6.44’N, 10°27.96’E). Animals from the

Wadden Sea were sampled at low tide. Epifaunal species were collected by forceps or

hand, and infaunal species by digging up the sediment with a spade to a depth of

approximately 25 cm and searching it by hand. Animals were placed in beakers filled

with a layer of wet sediment from the sampling site until further processing in the

laboratory. Sampling in Aarhus Bay was carried out from a research vessel by dredging

the sediment with a triangle net. Some animals such as shore crabs and ascidians were

sampled from rocks or pontoons in the harbor area of Aarhus. Sampled animals were

kept in buckets filled with seawater from the upper water column (15°C) until

incubation in the laboratory was started. The temperature of the water was measured at

each sampling site, and water samples were filtered (0.2 μm) and stored at -20°C until

nitrate concentration was measured using the VCl3 reduction method (Braman and

Hendrix 1989) with a chemiluminescence detector (CLD 66 S NO/NOx-Analyser, Eco

Physics).

Classification of species

The screening included Crustacea, Mollusca, Echinodermata, Polychaeta, and Ascidia

(Table 1). For each species, the affiliation to a feeding type and to a benthic habitat was

determined (Table 1). Species that feed by several feeding modes were assigned to their

dominant feeding mode. The description ‘infaunal + epifaunal’ refers to infaunal

species that feed at the sediment surface or in the water column. Species were further

characterized by their wet weight and by the presence/absence of a visible microbial

biofilm on exoskeletal surfaces such as molluscan shells, crustacean exoskeletons and

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shell plates of polychaetes (Table 1). Most species with sturdy external surfaces carried

such exoskeletal biofilms, but some of the crustacean and molluscan species (i.e.

Corophium volutator, Pagurus bernhardus, and Litopenaeus vannamei, Macoma

balthica, Scrobicularia plana, Cerastoderma edule) did not.

Table 1. List of taxa tested for N2O emission with sampling details (temperature and nitrate concentration in the overlying water column at the sampling site). Taxa are sorted by descending weight within each taxonomic group. Sampling sites—AB: Aarhus Bay; WS: Wadden Sea; AQ: aquaculture; Feeding types—C: carnivore; DF: deposit-feeder; FF: filter-feeder; G: grazer. Habitat—E: epifaunal; I: infaunal; EI: epifaunal + infaunal

Species Site Temp.

(°C)

Nitrate

(μM)

Wet weight

(g)

Feeding

type

Habitat Exoskeletal

biofilm

Ascidia Ascidia sp. AB 16 0-4 7.18 FF E Yes

Crustacea Carcinus maenas AB 15 0-4 2.95 C E Yes Pagurus bernhardus AB 7 0-4 2.81 C E No Corophium volutator WS 8 20 0.01 DF EI No

Echinodermata Echinocyamus pusillus AB 7 0-4 0.71 DF I No Echinocardium cordatum AB 7 0-4 0.27 DF I No

Mollusca Scrobicularia plana WS 15 20 4.63 DF EI No Cerastoderma edule WS 15 20 2.07 FF EI No Mytilus edulis AB 7 0-4 0.97 FF E Yes Macoma balthica WS 15 20 0.31 DF EI No Polyplacophoraa AB 7 0-4 0.27 G E Yes Littorina littorea WS 22 20 2.22 G E Yes Hinia reticulata AB 7 0-4 1.70 C EI Yes Gibbula sp. AB 7 0-4 0.78 G E Yes Hydrobia ulvae WS 21 20 0.01 G E Yes

Polychaeta Arenicola marina WS 8 20 2.06 DF I No Lepidonotus squamatus AB 7 0-4 0.49 C E Yes Nephtys hombergii WS 8 20 0.33 C I No Nereis diversicolor WS 8 20 0.15 DF EI No

Crustacea Litopenaeus vannamei AQ 28-30 1000 21.16 DF E No

a Not determined to genus level

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Rate of N2O emission

N2O emission of the specimens was determined by incubating freshly collected, living

animals (exception: Litopenaues vannamei) in gas-tight vials with septa that allowed

repeated sampling of the headspace for N2O. The incubations were standardized

regarding the environmental variables temperature (21°C) and oxygen (initially oxic

headspace), since the main goal of the screening was to search for species-specific

rather than environmental controls of the N2O emission potential. In many cases, the

standardized conditions in the incubation vial were different from those in the natural

habitat of the animals. Therefore, the N2O emission rates measured with this approach

represent potential rather than actual or in situ rates.

Incubation of animals was started after sampling, transport, and preparation of

incubation vials which took 3 to 5 h. Species were incubated in 3, 6, 10 or 100 ml

sterile, gas-tight vials, depending on their size and number of individuals. Most species

were found in sufficient quantity to prepare several vials per species with different

numbers of individuals (Table 2). Bivalves and ascidians were submerged in seawater to

allow the individuals to be active and thereby exchange gases with the incubation vial.

To the other species only a small volume of seawater was added (0.05-2 ml) to maintain

a moist atmosphere in the vials. Species from Aarhus Bay were supplied with 0.2 μm

filtered seawater collected while sampling the animals; species from the intertidal flat

were supplied with autoclaved seawater from the same site, collected during high tide

and stored in an opaque tank until used for incubations. Animals were thus exposed to

in situ nitrate and ammonium concentrations. The ammonium concentration in the

incubation vials was initially below the detection limit of 0.5 μM and may have

increased due to excretion of ammonium by the animals, which was in the range of

0.1�1.0 μmol individual�1 h�1 (Heisterkamp unpublished results). The aquacultured

shrimp Litopeaneus vannamei were killed in ice-water before incubating them in 100 ml

bottles with 2 ml of 0.2 μm filtered aquarium water that contained 1mM nitrate and 14

μM ammonium. Additionally, dissected guts of L. vannamei were incubated in 3 ml

exetainers (Labco) supplied with 50 μl of 0.2 μm filtered aquarium water.

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Table 2: Potential N2O emission rates in nmol ind.�1 h�1 and nmol g�1 h�1 of the 20 tested species (mean values). Number of replicates per species and range of individual numbers per incubation vial, as well as the initial N2O concentration and final N2O concentration in the incubation vials are listed (mean values).

N2O conc. (nM)

Species

N2O emission

(nmol g– 1 h– 1) N (range) Initial Highest

Ascidia sp. 0.043 ± 0.024 5(1–4) 5.9 54 Carcinus maenas 0.369 ± 0.137 3(1–3) 12.5 311 Pagurus bernhardus 0.020 ± 0.018 5(1–3) 8.5 167 Corophium volutator 0.955 ± 0.664 2(6–7) 10.2 123 Echinocyamus pusillus 0.040 ± 0.027 3(1–3) 12.7 40 Echinocardium cordatum 0.069 1 (5) 12.2 20 Scrobicularia plana 0.302 ± 0.083 3(2–3) 9.2 263 Cerastoderma edule 0.126 1 (5) 9.5 187 Mytilus edulis 0.269 ± 0.280 7 (1) 10.2 264 Macoma balthica 1.098 ± 1.066 7 (4–30) 9.8 287 Polyplacophoraa 0.471 ± 0.237 2 (6) 12.5 465 Littorina littorea 0.237 ± 0.208 6 (5–15) 9.7 167 Hinia reticulata 0.608 ± 0.265 7(1–3) 13.1 542 Gibbula sp. 0.107 ± 0.037 2(2–4) 13.1 345 Hydrobia ulvae 5.449 ± 1.822 4 (25 – 50) 10.7 463 Arenicola marina 0.045 ± 0.032 3 (1–2) 11.3 55 Lepidonotus squamatus 0.666 1 (3) 12.5 466 Nephtys hombergii 0.082 ± 0.053 3 (1–2) 0.1 5.6 Nereis diversicolor 0.398 ± 0.319 9 (1–2) 11.7 21 Litopenaeus vannamei 0.183 ± 0.066 6 (1) 12.5 250 aNot determined to genus level

Animals were cleaned from loosely attached sediment and algal tufts by washing them

in autoclaved seawater and drying them on paper tissue; the tightly attached biofilms

largely remained on the external surface of the animals. To explicitly test for effects of

this exoskeletal biofilm on the N2O emission potential, the snail Hinia reticulata was

incubated both with biofilm-covered shells and with shells that were cleaned by

thoroughly brushing them with a sterile toothbrush, although cleaning still left residues

of biofilm in the grooves of the shell surface.

The accumulation of N2O in the incubation vial was followed over a period of 4 to 6 h

by regularly taking gas samples and analyzing them by gas chromatography. Samples

from the Wadden Sea were measured with the GC 7890 (Agilent Technologies) with a

CP-PoraPLOT Q column, and samples from Aarhus Bay with the GC-8A (Shimadzu)

with a Porapak Q column. Both gas chromatographs were equipped with a 63Ni electron

capture detector. Injection volumes were 1 ml for the samples analyzed with the GC

7890, and 0.3 ml for samples analyzed with the GC-8A. After each headspace sampling,

72


the incubation vials were pressure-equilibrated with air by inserting a hypodermic

needle through the septum for 1 s. On both GCs, calibration standards were prepared by

adding known amounts of N2O to N2-flushed gas-tight bottles of known volume and

analyzed repeatedly during the incubation. The linear part of the increase of the N2O

concentration in the incubation vials over time was used to calculate the potential N2O

emission rate per individual and per biomass. The dilution of the gas phase and the

equilibrated distribution of N2O between the gas and water phases (Weiss and Price

1980) were taken into account when calculating the potential N2O emission rate. This

rate corresponds to the net N2O production rate (i.e., gross production less consumption)

and thus also depends on N2O levels. Since the N2O reduction rate was not directly

assessed, the initial and final N2O concentrations in the incubation vials are reported in

Table 2 so that the experiments can be reproduced.

Rate of total denitrification

To determine the potential rate of total denitrification (i.e., production of N2+N2O) in

the shrimp gut, freshly killed Litopenaues vannamei were dissected and the guts were

incubated in an atmosphere of 10% acetylene and 90% dinitrogen gas. Acetylene

inhibits the last step of denitrification (Sørensen 1978) and thus the accumulation of

N2O in the incubation vials is indicative of total denitrification. The linear increase of

N2O concentration in the incubation vials over time was used to calculate the potential

total denitrification rate per gut.

Statistical analysis

The potential N2O emission rates were tested for correlation with the species traits

Feeding type, Habitat, Exoskeletal biofilm, and Weight using the statistical analysis

software SPSS. The categories within the species traits Feeding type, Habitat, and

Exoskeletal biofilm were ranked according to their hypothesized effects on N2O

emission rates and were transformed into a numerical code that could be used for

correlation analysis (Table 3). The hypotheses were that the rate of N2O emission is

positively correlated to (1) the amount of ingested bacteria, (2) the availability of

nitrate, and (3) the presence of a microbial biofilm growing on external surfaces of the

animal. The ranking of the categories was based on the assumptions that (1) the amount

73


of ingested bacteria is determined by the feeding type and increases from carnivores

over grazers and deposit-feeders to filter-feeders; (2) the nitrate concentration varies

with habitat, being highest in the water column and lowest in the sediment; and (3) shell

and exoskeleton provide colonization surfaces for microbial biofilms. The high rank of

filter-feeders regarding the amount of ingested bacteria may be questioned because only

few bivalve species filter unattached bacteria (e.g., Mytilus edulis, McHenery and

Birkbeck 1985). However, species that filter-feed close to the sediment surface, where

the concentration of suspended detritus is particularly high, ingest large amounts of

attached bacteria (Kach and Ward 2008).

Table 3: Species traits and phenotypes used for statistical analysis of N2O emission by marine invertebrates. Phenotypes were sorted according to their hypothesized promotion of N2O production (Hypothesis) and then numerically coded (Value). Species trait Phenotype Hypothesis Value

Feeding type Carnivore (predator + scavenger) 0

Grazer 1

Deposit-feeder 2

Filter-feeder

Increasing

number of N2O-

producing gut

bacteria 3

Habitat Infaunal 0

Infaunal + epifaunal 1

Epifaunal

Increasing nitrate

availability

2

Exoskeletal biofilm No 0

Yes

More biofilm

bacteria 1

Results

The potential N2O emission rates of coastal marine invertebrate species ranged

from 0 to 1.354 nmol ind.�1 h�1 (Figure 1, Table 2) with an average rate of

0.320 nmol ind.�1 h�1. The weight-specific emission rates ranged from 0 to 0.598 nmol

g�1 h�1 with an average rate of 598 nmol g�1 h�1 (Table 2). The highest potential N2O

emission rate of 3.569 nmol ind.�1 h�1 was found for the aquacultured shrimp

Litopenaeus vannamei (not included in the above rates) that is exposed to very high

74


nitrate concentrations (� 1 mM) and to high temperatures (28-30°C) in the rearing tanks

(Table 1). The N2O emission rate of dissected guts of L. vannamei was almost as high

as the N2O emission rate of the whole animal (Figure 2). Dissected guts showed a total

denitrification rate of 12 nmol ind.�1 h�1 under anoxic conditions (Figure 2).

N2O (nmol ind.-1 h-1)

0,00 0,25 0,50 0,75 1,00 1,25 1,50

Ascidia sp.

Carcinus maenasPagurus bernhardusCorophium volutator

Echinocyamus pusillusEchinocardium cordatum

Scrobicularia plana

Macoma balthicaMytilus edulis

Cerastoderma edule

Polyplacophora

Hinia reticulataLittorina littorea

Gibbula sp.

Hydrobia ulvae

Lepidonotus squamatusNephtys hombergii

Arenicola marina

Nereis diversicolorexoskeletal biofilm

no exoskeletal biofilm

0,0 0,5 1,0 1,5 2,0 2,5 3,0 3,5 4,0

Litopenaeus vannamei

Aquaculture

Coastal marine ecosystem

Figure 1: Potential N2O emission rates of various marine invertebrate taxa. Individuals were incubated in gas-tight vials under oxic conditions at 21°C and N2O emission was analysed by GC measurements over 4-6 h. Species are grouped taxonomically and within each taxonomic group, species are sorted by descending weight. For species with at least 3 replicates analysed, mean rates + s.d. are shown.

75


Dissected gut

N2O

(n

mo

l in

d.-1

h-1

)

0

2

4

6

8

10

12

14

16

Dissected gut(-oxygen +acetylene)

Whole animal

Figure 2: Potential N2O emission by the shrimp Litopenaeus vannamei and its dissected guts under oxic conditions at 21°C. Dissected guts of L. vannamei were also incubated under anoxic conditions with 10% acetylene that inhibits the last step of denitrification. The resulting N2O produc-tion indicates total denitri-fication. Means + s.d. are shown (n=3-6).

The nitrate concentrations at the sampling sites in the Wadden Sea and Aarhus Bay

were low (0-20 μM), and temperature was 7-8°C (exception: 15-22°C at the Wadden

Sea site in May 2008; Table 1). The capacity to emit N2O occurred across all taxonomic

groups and was not restricted to a certain feeding type (Table 1). Most species

possessing a shell or exoskeleton had potential N2O emission rates higher than the

average rates (e.g., the common periwinkle Littorina littorea and the shore crab

Carcinus maenas). These conclusions were also true when the rate of N2O emission was

expressed per gram body weight (Table 2). The potential N2O emission rates per

individual tended to be higher for larger species than for smaller species (e.g. the

bivalves Scrobicularia plana vs. Macoma balthica), while the highest potential N2O

emission rates per gram body weight were shown by the smallest species (e.g. Hydrobia

ulvae, Corophium volutator).

The correlation analysis revealed that the potential N2O emission rate per individual was

significantly positively correlated with the body weight with a Pearson coefficient of R

= 0.506 (p = 0.027) for linear correlation and with a Spearman coefficient of R = 0.728

(p < 0.001) for non-linear correlation. The species-traits Habitat and Exoskeletal biofilm

showed significant positive non-linear correlations with the potential N2O emission rate

per individual with Spearman coefficients of R = 0.460 (p = 0.047) and R = 0.481

(p = 0.037), respectively. No correlation between the potential N2O emission rate and

the feeding type was found (Spearman coefficient of R = -0.135, p = 0.581).

76


The importance of the species trait Exoskeletal biofilm was further highlighted by the

comparison of the N2O emission rates of the snail Hinia reticulate, which was measured

both with the natural biofilm on the surface of the shell and with cleaned shell surfaces.

The snails with an exoskeletal biofilm emitted more N2O than the cleaned individuals

during the incubation period of 4.5 h (Figure 3). The mean potential N2O emission rate

of the biofilm-covered individuals (1.108 nmol ind.�1 h�1) was about 3.5 times higher

than the rate of the cleaned individuals (0.306 nmol ind.�1 h�1). The mean potential N2O

emission rate of biofilm-covered and cleaned individuals were assessed by a t-test and

marginally failed significance with p = 0.057 (T = -3.06 and df = 2.92).

Time [h]

0 1 2 3 4 5

N2O

(n

mo

l in

d.-1

)

0

1

2

3

4

5

6

7cleaned

biofilm-covered

Figure 3: Potential N2O emission by cleaned and biofilm-covered individuals of the snail Hinia reticulata during the incubation period of 4.5 h. Means + s.d. are shown (n=3).

Discussion

The present study revealed that many coastal marine invertebrate species emit N2O,

representing a source that have been overlooked. The average potential N2O emission

rate of 19 marine invertebrate species was 0.320 nmol ind.�1 h�1, excluding the

aquacultured shrimp Litopenaeus vannamei, which had an exceptionally high rate. For

20 freshwater invertebrate species, an average potential N2O emission rate of only

0.072 nmol ind.�1 h�1 was reported (Stief et al. 2009). In addition to the higher average

rate, the N2O emission potential of marine invertebrates is apparently influenced by

species-specific traits (i.e., body weight, habitat, and presence of an exoskeletal biofilm)

77


that differ from those that influence the N2O emission potential of freshwater species

(i.e., feeding type) (Stief et al. 2009).

Correlation with species traits

At a first glance, the positive correlation with the body weight suggests that larger

animals with presumably larger guts produce more N2O than smaller animals because of

the larger number of microbes passing their gut. This interpretation is consistent with

the hypothesis that, in marine invertebrates, N2O production is also mediated by

ingested microbes, as is the case for earthworms and freshwater invertebrates (Drake et

al. 2006, Stief et al. 2009). The correlation between potential N2O emission rate and the

presence of an exoskeletal biofilm suggests that N2O production associated with marine

invertebrates is not always due to denitrification in the gut (as proven for the

aquacultured shrimp Litopenaeus vannamei), but may also result from microbial activity

on external surfaces of the animal. Lower potential N2O emission rates of individuals of

the snail Hinia reticulata with an experimentally cleaned shell surface further

substantiate that N2O production is also linked to microbial activities in the exoskeletal

biofilm. Furthermore, for this type of animal-associated N2O production, the shell of

larger animals is presumably colonized with larger numbers of bacteria involved in N2O

production, which is in line with the weight-dependence of N2O emission. The

microbial pathway for biofilm-associated N2O production still needs to be identified.

Depending on the oxygen availability inside the biofilm, nitrification or denitrification

or both might contribute to the production of N2O (Meyer et al. 2008). Likewise, N2O

production in the exoskeletal biofilm might be driven by ammonium from animal

excretion or by nitrate from the water column, or by both. If an oxic-anoxic transition

zone prevails in the biofilm, then nitrification and denitrification are probably coupled,

as known for sediments in which denitrification is driven by nitrate from nitrification

(Jenkins and Kemp 1984). Thick biofilms were not established on the exoskeleton of

every molluscan and crustacean species tested in the present study. The exoskeleton of

Corophium volutator, Pagurus bernhardus, and Litopenaeus vannamei, for instance,

may not allow the formation of a persistent biofilm due to rather short time intervals

between molting events, and the shells of infaunal molluscs (i.e., Macoma balthica,

Cerastoderma edule and Scrobicularia plana) may not be suitable for the formation of

an exoskeletal biofilm due to physical abrasion in the sediment. It remains to be

78


investigated whether certain freshwater invertebrate species have persistent biofilms on

external surfaces of their body that produce N2O.

Habitat (a proxy for the nitrate availability in the immediate environment of the animal)

was also significantly correlated with the N2O emission rate. The high potential

emission rate of the epifaunal shrimp Litopenaeus vannamei, which is exposed to very

high nitrate concentrations, agrees with this assumption. The effect of the habitat on

N2O emission could be greater during autumn and winter, when nitrate concentrations

in the water column at the 2 study sites are higher than in spring and summer

(Kieskamp et al. 1991, Sømod 2005) and most of the studied animals are also abundant

and active. Additionally, the habitat of marine macrofauna may have an influence on the

formation of exoskeletal biofilms. Epifaunal species are expected to carry thicker and

more persistent biofilms on their external surfaces than infaunal species because of

lower abrasion forces in the water column. This interaction of “habitat” and

“exoskeletal biofilm” is supported by our observations on, for instance, Littorina

littorea (epifaunal, visible biofilm, high N2O emission potential) vs. Macoma balthica

(infaunal, no visible biofilm, lower N2O emission potential).

N2O emission rate and species feeding type and diet were not correlated, which

contrasts with the finding that N2O emission of freshwater invertebrates is diet-

dependent (Stief et al. 2009). Since marine species are usually larger and have longer

guts and gut residence times than the freshwater species (Bayne et al. 1987, Navarro et

al. 1993), bacteria might be exposed long enough to anoxic conditions in the gut to

express the full set of denitrification genes. In that case, complete denitrification will

prevail and the main product will be dinitrogen rather than N2O. Conversely, many of

the ingested sediment bacteria might be efficiently digested in the gut of marine

detritivorous species due to a high lysozyme activity (Lucas and Bertru 1997, Plante

and Mayer 1994), which inhibits microbial N2O production. Lysozyme activity of

dissected guts was approximately 5 times higher for the ragworm Nereis diversicolor

(a marine non-emitter) than for the mayfly larva Ephemera danica (a freshwater

emitter) (P. Stief unpublished results).

79


Ecosystem perspective

Many species tested positive for N2O emission in the present study are very abundant in

coastal soft-bottom habitats; Macoma balthica and Cerastoderma edule can reach

densities of 1000 ind. m-2 (Fujii 2007), Scrobicularia plana 250 ind. m-2 (Cabral and

Murta 2004), and Arenicola marina 100 ind. m-2 (Flach and Beukema 1994). The mud

snail Hydrobia ulvae can reach densities of up to 100 000 ind. m-2 in intertidal

sediments (Barnes 1999). This epifaunal species emits N2O directly into the water

column or the atmosphere without diffusion through the sediment, as it lives at the

sediment surface where it can be exposed to high nitrate concentrations and

temperatures. Taking its potential N2O emission rate of 0.068 nmol ind.�1 h�1, this small

snail could emit 6.8 μmol N2O m-2 h�1, which is on the same order of magnitude as the

benthic N2O fluxes reported for estuarine intertidal sediments (Middelburg et al. 1995)

and intertidal rocky biofilms (Magalhaes et al. 2005). For infaunal species,

extrapolations are less robust because N2O conversion may also take place inside the

burrows of the animals (Stief and Schramm 2010). N2O produced by certain infaunal

species might be partially consumed while diffusing towards the sediment surface

(Meyer et al. 2008), whereas other infaunal species could increase the benthic N2O flux

much more by their bioirrigation activity than by stimulating N2O production in their

gut or in exoskeletal biofilms (Stief and Schramm 2010). A second difficulty in scaling

up animal-associated N2O production to ecosystem level lies in the discrepancy between

potential and in situ rates. The contribution of animal-associated N2O production to

overall benthic N2O emission can be better estimated from rate measurements made at

different times of the year at the prevailing environmental conditions (Stief et al. 2010,

Stief and Schramm 2010). A rather constant N2O emission rate can be expected for the

aquacultured species Litopenaeus vannamei, since it is exposed to the same conditions

throughout the year. Given its very high potential N2O emission rate and the high

growth rates of the aquaculture industry, N2O emission by other aquacultured species

should be investigated.

Conceptually, N2O production associated with marine and freshwater invertebrates

constitutes a link between reactive nitrogen (i.e., nitrate and ammonium) in aquatic

ecosystems and N2O in the atmosphere that has been overlooked. Aquatic invertebrates

complement the known sites of N2O production in the sediment with 3 additional

80


microsites of N2O production: 1) the anoxic gut, a transient microbial habitat in which

denitrification prevails (Stief et al. 2009); 2) the burrow, a microbial habitat with

fluctuating conditions in which nitrification and denitrification co-occur (Svensson

1998); and 3) the exoskeletal biofilm, a microbial habitat with a yet unknown

microenvironment in which nitrification and/or denitrification might occur (present

study). The environmental controls of sedimentary and animal-associated N2O

production might be similar (e.g. higher N2O production rates at higher temperature and

nitrate or ammonium concentrations) and require further investigation.

Acknowledgements

We thank Hans Brix for providing the gas chromatograph. We are grateful for the help

and assistance of Torben Vang and Leif Flensborg, the crew of the research vessel

‘Genetica II’ (Aarhus University). This research was supported by the Max Planck

Society, the Danish Research Council and the German Science Foundation (STI202/6).

81


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84

Chapter 3

Marine mollusc species that were investigated for nitrous oxide production in shell biofilms

(a) Mytilus edulis (Blue Mussel) (b) Littorina littorea (Common Periwinkle)

(c) Hinia reticulata (Netted Dog Welk)

85


processes, precursors and relative importance

Ines M. Heisterkamp1, Andreas Schramm2, Lone H. Larsen2,

Nanna B. Svenningsen2, Gaute Lavik1, Dirk de Beer1, Peter Stief1


Celsiusstraße 1, 28359 Bremen, Germany

2Department of Bioscience, Microbiology, Aarhus University,


Accepted in Environmental Microbiology

87

Chapter 3 N2O production in shell biofilms

Abstract

Emission of the greenhouse gas nitrous oxide (N2O) from freshwater and terrestrial

invertebrates has exclusively been ascribed to N2O production by ingested denitrifying

bacteria in the anoxic gut of the animals. Our study of marine molluscs now shows that

also microbial biofilms on shell surfaces are important sites of N2O production. The

shell biofilms of Mytilus edulis, Littorina littorea, and Hinia reticulata contributed

18-94% to the total animal-associated N2O emission. Nitrification and denitrification

were equally important sources of N2O in shell biofilms as revealed by 15N-stable

isotope experiments with dissected shells. Microsensor measurements confirmed that

both nitrification and denitrification can occur in shell biofilms due to a heterogeneous

oxygen distribution. Accordingly, ammonium, nitrite, and nitrate were important drivers

of N2O production in the shell biofilm of the three mollusc species. Ammonium

excretion by the animals was found to be sufficient to sustain N2O production in the

shell biofilm. Apparently, the animals provide a nutrient-enriched microenvironment

that stimulates growth and N2O production of the shell biofilm. This animal-induced

stimulation was demonstrated in a long-term microcosm experiment with the snail H.

reticulata, where shell biofilms exhibited the highest N2O emission rates when the

animal was still living inside the shell.

88


Introduction

Nitrous oxide (N2O) is a highly potent greenhouse gas that significantly contributes to

global warming (Forster et al., 2007) and to the destruction of the stratospheric ozone

layer (Ravishankara et al., 2009). This atmospheric trace gas is mainly produced in soils,

sediments and water bodies by nitrifying and denitrifying bacteria (Mosier et al., 1998;

Stein and Yung, 2003). Ammonia-oxidizing bacteria and archaea (AOB and AOA)

produce N2O during the oxidation of NH4+ to NO2

− the first step of nitrification (Goreau

et al., 1980; Santoro et al., 2011), and during “nitrifier denitrification”, a process in

which NH4+ is oxidized to NO2

− that is subsequently reduced to NO, N2O and

sometimes N2 (Ritchie and Nicholas, 1972; Poth and Focht, 1985; Bock et al., 1995).

Heterotrophic denitrifying bacteria produce N2O as an intermediate when reducing

NO3− or NO2

− via NO and N2O to N2 under suboxic or anoxic conditions (Zumft, 1997).

High N2O emission rates have been reported for earthworms and several freshwater and

marine invertebrates (Karsten and Drake, 1997; Stief et al., 2009; Heisterkamp et al.,

2010). The N2O production associated with these invertebrates has been attributed to

incomplete denitrification in the gut of the animals. The gut microenvironment of N2O-

emitting animals is characterised by anoxia and the availability of nitrate and labile

organic carbon sources, and thus stimulates the denitrification activity of ingested

microorganisms (Horn et al., 2003; Stief and Eller, 2006). Accordingly, N2O emission

from freshwater invertebrates largely depends on the amount of ingested bacteria, which

varies with the feeding-type of the animal (Stief et al., 2009), and on the nitrate

concentration (Stief et al., 2010; Stief and Schramm, 2010). For marine invertebrates,

however, N2O emission rate and feeding type were not correlated (Heisterkamp et al.,

2010). Instead, N2O emission rate was marginally correlated with the presence of a

microbial biofilm on external surfaces of the animals (Heisterkamp et al., 2010). These

findings suggested that N2O emission by marine invertebrates may not only be due to

N2O production by denitrifying bacteria in the gut, but also due to N2O production in

microbial biofilms on the shell or exoskeleton of the animals.

89


The objectives of this study were therefore to (i) quantify the contribution of the shell

biofilm to total N2O emission from marine molluscs, (ii) quantify the contribution of

nitrification and denitrification to N2O production in shell biofilms, and (iii) assess the

effect of the animal itself on N2O production in shell biofilms. To this end, short-term

measurements with freshly collected specimens, stable isotope incubations of dissected

shells, and long-term sediment-microcosm experiments were conducted with three

marine mollusc species, Mytilus edulis (Blue Mussel), Littorina littorea (Common

Periwinkle), and Hinia reticulata (Netted Dog Welk). These molluscs typically possess

a shell biofilm and are common in marine coastal environments of the North Atlantic

and Baltic Sea.

Material and Methods

Sampling of animals

We collected three marine mollusc species that inhabit different habitats in the North

Atlantic and Baltic Sea and exhibit distinct feeding-modes (Table 1). The snail Littorina

littorea was sampled from a mixed intertidal flat near Dorum-Neufeld in the German

Wadden Sea (N53° 44' 3.733" E8° 30' 24.201"). The mussel Mytilus edulis and the snail

Hinia reticulata were collected in Aarhus Bay, Denmark. M. edulis was taken from the

mole of Marselisborg Harbor in Aarhus (N56° 8' 16.875" E10° 13' 3.945") and

H. reticulata from a very shallow part of Aarhus Bay close to Skaering (N56° 13'

39.045" E10° 18' 51.154"). Animals were kept in buckets filled with aerated seawater

and a thin layer of sediment from the sampling site (M. edulis only in seawater) until

used for experiments in the laboratory. All experiments were performed within 2 days

after sampling.

90


Table 1: Characteristics of the three mollusc species. Wet weight of the whole animal and dissected shell, as well as NH4

+ excretion rate of the animal are listed as means (± SD), n = 4-62.

Mytilus edulis (Blue Mussel)

Littorina littorea (Common Periwinkle)

Hinia reticulata (Netted Dog Whelk)

Taxonomic class Bivalvia Gastropoda Gastropoda

Feeding type Filter-feeder Grazer Scavenger

Habitat Intertidal rocks Intertidal sediments & rocks

Sublittoral sandy sediments

Weight of whole animal (g) 20.7 (1.8) 1.9 (0.1) 1.5 (0.7)

Weight of dissected shell (g) 13.7 (2.9) 1.4 (0.1) 0.9 (0.3)

NH4+ excretion rate

(nmol g−1 h−1) 64 (36) 27 (13) 154 (110)

NH4+ excretion rate

(nmol ind−1 h−1) 969 (549) 69 (16) 271 (191)

Nitrous oxide emission rates of whole animals and their dissected shells

N2O emission rates of freshly collected, living animals and of the animals’ shell were

measured by incubating whole animals or dissected shells in gas-tight vials and

following N2O production over 4 h. Shells were dissected from living animals by

cracking the shell and carefully removing the animal tissue with ethanol-rinsed forceps

and scalpels taking care not to damage the shell biofilm. Artificial seawater was

prepared by dissolving Red Sea salt (Red Sea Fish Farm Ltd. Eilat, Israel) in autoclaved

MilliQ water to the in situ salinity of 22 psu, aerated over night, adjusted to pH 8.2, and

amended with 50 μM NH4+ and 50 μM NO3

−. Ammonium concentration was thus

higher than the typical water column concentration, but nitrate concentration was in the

range of naturally occurring concentrations (van Beusekom and de Jonge, 2002; van

Beusekom et al., 2008). Whole animals and dissected shells of M. edulis and L. littorea

were incubated in 100-ml Duran bottles filled with 50 ml and 30 ml of artificial

seawater, respectively, and sealed with rubber stoppers. Whole animals and dissected

shells of H. reticulata were incubated in gas-tight 12-ml Exetainer vials (Labco, High

Wycombe, UK), to which 3 ml of seawater was added. The remaining headspace

consisted in all cases of atmospheric air. Additionally, dissected shells of the three

91


species were incubated in artificial seawater that contained 2% of ZnCl2 (50% w/v) to

inhibit biological activity. For each species, 3-8 replicates were run with either

1-2 individuals or dissected shells per incubation vial (M. edulis and H. reticulata) or

5-8 individuals and 8-11 dissected shells per incubation vial (L. littorea). Vials were

placed on a shaker during incubation to enforce the equilibration of N2O between water

and headspace and were shaded from direct daylight (light intensity approximately

5 μmol photons m−2 s−1). All incubations were made in a temperature-controlled

laboratory at 21°C. Analysis of N2O production by gas chromatography (GC 7890

Agilent Technologies) and calculation of N2O emission rates were performed as

described in Heisterkamp et al. (2010).

15N-stable isotope experiments with dissected shells

For each species, dissected shells were incubated in three different 15N-tracer treatments

that were designed to specifically ascribe the production of the double-labelled 46N2O (15N15NO) to nitrification, denitrification of nitrate and denitrification of nitrite

(Table 2). To be able to distinguish between N2O production by nitrification and N2O

production by coupled nitrification-denitrification or nitrifier denitrification, a ten times

higher 14NO2− than 15NH4

+ concentration was added to the nitrification treatment. Any 15NO2

− generated from nitrification is thus diluted by the high 14NO2− concentration,

which makes the combination of two 15NO2− and subsequent 46N2O production via

coupled nitrification-denitrification or nitrifier denitrification very unlikely. The double-

labelled 46N2O can thus be ascribed to the combination of two 15NH4+ and is indicative

of nitrification. The single-labelled 45N2O (15N14NO) could have been produced by

random pairing of 15NH4+ with unlabelled 14NH4

+ that were present or produced in the

biofilm, or by coupled nitrification-denitrification and nitrifier denitrification that

combine one 15N from 15NH4+ with one 14N from 14NO3

− or 14NO2− to form 45N2O. In

the denitrification treatment, any 46N2O production must have exclusively resulted from

the combination of two 15NO3− ions. 45N2O was produced by random pairing of 15NO3

−

and 14NO3− or 14NO2

− that were present or produced in the biofilm.

92


Tab

le 2

: Sta

ble

isot

ope

incu

batio

ns m

ade

with

dis

sect

ed sh

ells

of M

. edu

lis,L

. litt

orea

and

H. r

etic

ulat

a. L

iste

d ar

e th

e co

ncen

tratio

ns o

f 15N

and

14N

that

w

ere

adde

d to

eac

h tre

atm

ent,

the

requ

ired

com

bina

tions

of 14

N a

nd 15

N to

form

45N

2O a

nd 46

N2O

, and

the

prob

abili

ties

of th

e fo

rmat

ion

of 44

N2O

, 45N

2O

and

46N

2O a

ssum

ing

rand

om i

soto

pe p

airin

g of

the

uni

form

ly m

ixed

14N

and

15N

spe

cies

. 14N

Ox- i

nclu

des

14N

O2- a

nd 14

NO

3- tha

t w

ere

adde

d to

the

in

cuba

tions

, alre

ady

pres

ent o

r pro

duce

d in

the

shel

l bio

film

s. In

the

nitri

ficat

ion

treat

men

t, 45 N

2O c

ould

als

o ha

ve b

een

prod

uced

by

the

com

bina

tion

of

labe

lled

15N

H4+ a

nd u

nlab

elle

d 14

NH

4+ that

was

alre

ady

pres

ent o

r pro

duce

d in

the

shel

l bio

film

s.

Ran

dom

isot

ope

pair

ing

[500

×14N

O2− ]2 +

2[5

00×14

NO

2− ][50

×15N

H4+ ] +

[50×

15N

H4+ ]2

[50×

15N

O3− ]2

[500

×14N

O3− ]2 +

2[5

00×14

NO

3− ][50

×15N

O2− ] +

[50×

15N

O2− ]2

46N

2O

15N

H4+

+ 15

NH

4+

15N

O3− +

15N

O3−

15N

O2− +

15N

O2−

45N

2O

14N

H4+ /14

NO

x− + 15

NH

4+

14N

Ox− +

15N

O3−

14N

Ox− +

15N

O2−

14N

add

ed

14N

O2− (5

00 μ

M)

−

14N

O3− (5

00 μ

M)

15N

trac

er

15N

H4+ (5

0 μM

)

15N

O3− (5

0 μM

)

15N

O2− (5

0 μM

)

Tre

atm

ent

Nitr

ifica

tion

Den

itrifi

catio

n of

nitr

ate

Den

itrifi

catio

n of

nitr

ite

93


In the 15NO2− treatment, a ten-fold higher 14NO3

− than 15NO2− concentration was added

to dilute the 15NO3− generated from nitrification and thereby make random pairing of

two 15NO3− and subsequent production of 46N2O via denitrification of nitrate very

unlikely. The 46N2O production can thus be ascribed to denitrification of nitrite. The

single-labelled 45N2O was produced by the combination of 15NO2− with 14NO3

− and/or 14NO2

− that were present or produced in the biofilms. Assuming random isotope pairing

of the uniformly mixed 14N and 15N species, the probabilities of the formation of 44N2O, 45N2O and 46N2O can be calculated as follows [14N]2 + 2[14N][15N] + [15N]2, in analogy

to the formula used by Nielsen (1992) to calculate the production of 28N2 from the 29N2

and 30N2 production ratio. The 14N to 15N ratio of 10 in the 15NH4+ and 15NO2

−

treatments, should thus result in a 44N2O:45N2O:46N2O ratio of 100:20:1 ([10 14N]2 +

2[10 14N][15N] + [1 15N]2).

Three replicates per species and treatment were set up by dissecting shells of freshly

sampled animals as described above and incubating them in 100-ml Duran bottles that

were filled with 100 ml autoclaved and aerated artificial seawater and sealed gas-tight

with rubber stoppers. The remaining bottle volume of 25 ml contained atmospheric air

to ensure that the oxygen concentration in the vial did not drop below 50% air-

saturation (tested by microsensor measurements after incubations). For L. littorea and

H. reticulata, 5-7 shells were incubated per vial, for M. edulis one individual shell per

vial. Wet weights of dissected shells were determined before starting the incubations.

The dissected shells were incubated on a shaker for 6 h at 21°C and low light conditions

(approximately 5 μmol photons m−2 s−1). At time points 0, 1, 2, 3, 4 and 6 h, water

samples (3 ml) were taken with a syringe and transferred to 6-ml He-flushed exetainers

prefilled with 0.1 ml saturated HgCl2. The water volume withdrawn from the incubation

vial was replaced with the respective 15N tracer solution to avoid under-pressure in the

vials. Each exetainer was spiked with 25 μl non-labelled N2O to be above the detection

limit of the mass spectrometer. The equilibrated headspace of the exetainer was then

analyzed for 44N2O, 45N2O and 46N2O concentrations by gas chromatography-isotope

ratio mass spectrometry (GC-IRMS; VG Optima, Manchester, UK). The excess

concentrations of 45N2O and 46N2O were calculated from the ratios 45N2O: 44N2O and 46N2O: 44N2O using a 100% N2O standard in analogy to the air standard used in the

isotope pairing technique of Nielsen (1992) where the excess of 29N2 or 30N2 is

determined from the 29N2: 28N2 and 30N2: 28N2 ratios, respectively. The linear increase of

94


excess 45N2O and 46N2O over time was used to calculate 45N2O and 46N2O net

production rates taking the dilution of the water phase and the equilibrated distribution

of N2O between the water and gas phases (Weiss and Price, 1980) into account.

Nitrous oxide production in shell biofilms in relation to nitrification and denitrification rates

The N2O production rates of dissected shells were measured under (a) oxic atmosphere

and addition of 50 μM NH4+ (nitrification assay), and (b) anoxic atmosphere and

addition of 50 μM NO3− (denitrification assay). Both assays were set up in 100-ml

Duran bottles filled with 20-30 ml aerated or He-flushed artificial seawater and were

sealed with rubber stoppers. The headspace of the oxic and anoxic incubation assays

was air and He gas, respectively. Analysis of N2O by gas chromatography and

calculation of N2O production rates were made as described above.

Potential ammonium oxidation rates of shells were measured by incubating freshly

dissected shells in 20 ml (60 ml for M. edulis) aerated artificial seawater containing

50 μM NH4+ and 20 mM NaClO3. The oxidation of nitrite to nitrate in nitrification is

inhibited by NaClO3 (Belser and Mays, 1980). Nitrite production is thus indicative of

ammonium oxidation, since the produced nitrite is not further oxidized to nitrate.

Additionally, a control without inhibitor was run. Incubation assays were aerated via a

hypodermic needle and stirred at 100 rpm on a multi-plate stirrer during the incubation

period of 4 h. Water samples (1 ml) were taken hourly and immediately frozen at -20°C

until nitrite concentration was measured using the NaI reduction method (Braman and

Hendrix, 1989) with a chemiluminescence detector (CLD 86 S NO/NOx- Analyser, Eco

Physics, Germany). The resulting nitrite production in the incubation vial was then used

to calculate the ammonium oxidation rate.

To determine the potential rate of total denitrification (i.e., production of N2 + N2O),

dissected shells were incubated in 100-ml Duran bottles filled with 20-30 ml anoxic

artificial seawater adjusted to 50 μM NO3−. The bottles were sealed with rubber

stoppers and the headspace was purged with He gas. One tenth of the headspace was

replaced by acetylene which inhibits the last step of denitrification (Sørensen, 1978) and

95


thus the accumulation of N2O in the incubation vials indicates total denitrification. A

control without acetylene was also run.

For each of the three species and four assays, three replicates with 1-5 shells per vial

were set up. All incubations were made at 21°C. The N2O yields from nitrification and

denitrification were calculated as the percentage of N2O/NO2− and N2O/N2+N2O

expressed per mol N.

Oxygen concentration in shell biofilms

Oxygen concentrations in the shell biofilms were measured with an oxygen microsensor

(Revsbech, 1989). The microsensor was calibrated in artificial seawater (19°C, salinity

of 22 psu) at 0 and 100% air-saturation by purging with nitrogen gas and synthetic air,

respectively. Small pieces of dissected shells were placed in a flow-cell which was

continuously flushed with oxygenated artificial seawater (Stief and Eller, 2006). Aided

by a dissection microscope, the microsensor tip was positioned at the shell surface and

vertical steady-state concentration profiles were recorded through the biofilm and the

diffusion boundary layer to a distance of maximally 2.4 mm from the shell surface in

increments of 0.0175 mm or 0.035 mm. Profiles were run at randomly chosen positions

during light (1220 μmol photons m−2 s−1) and dark conditions.

Effect of the animal on N2O production in shell biofilms

A long-term sediment-microcosm experiment with H. reticulata was set up with four

treatments: a) animals covered with the natural shell biofilm (A+), b) animals with the

shell biofilm removed with sterile scalpels and ethanol (A-), c) intact shells covered

with the natural biofilm but the animal removed (S+), and d) intact shells with the

biofilm and the animal removed (S-). The experiment was conducted in four

recirculating flow-through aquaria (30 cm long × 20 cm wide × 10 cm high) each of

which was filled with sediment and some seaweed from the sampling site and

continuously supplied from its own 50 L original seawater reservoir for a period of

53 days. The microcosms were illuminated from above (40 μmol photons m−2 s−1 light

intensity at the sediment surface) at a 16 h light to 8 h dark cycle throughout the whole

experiment.

96


The N2O production of dissected shells was investigated on day 1, 20, 33, and 53 of the

experiment. In treatments A+ and A-, animals were removed from the shells as

described above. Four dissected shells per treatment and sampling day were incubated

in 6-ml exetainers filled with 1 ml of 0.2-μm filtered seawater from the respective

microcosm. N2O production was followed over 4 h by taking headspace samples hourly

and analysing them as described above.

After the N2O measurements, dissected shells were immediately frozen at -20°C for

later analysis of the protein content of the biofilms (see below). Water samples were

taken from the four flow-through microcosms throughout the experiment and stored at

-20°C until ammonium concentration was analysed photometrically (Bower and Holm-

Hansen, 1980), and nitrate and nitrite concentrations by the VCl3 and NaI reduction

method, respectively (Braman and Hendrix, 1989). Correlation analysis of N2O

production rates and protein contents of dissected shells as well as DIN concentrations

in the water column of the microcosms were made with the statistical analysis software

SPSS (SPSS Inc., U.S.A.).

Protein content of shell biofilm

Dissected shells were thawed and incubated in 5 ml 0.5 N NaOH in a water bath at

80°C for 20 min to extract the proteins. The supernatant was collected after

centrifugation at 3000 rpm (1750 g) for 5 min. This extraction procedure was repeated

three times. The supernatants from the three extractions were then analysed by the

Lowry protein assay (Lowry et al., 1951).

Ammonium excretion rates

Freshly collected whole animals were weighed and incubated for 3 h in artificial

seawater continuously mixed by a magnetic stirrer. Per species, 6 incubations

containing from 1 individual (M. edulis) to up to 6 individuals (L. littorea) were set up.

Water samples were taken every 0.5 h and frozen at -20°C. Accumulation of ammonium

in the water samples was measured by spectrophotometry (Bower and Holm-Hansen,

1980).

97


Results

Nitrous oxide emission rates of whole animals and their dissected shells

N2O was produced during incubation of both whole animals and biofilm-covered

dissected shells (Fig. 1). The N2O emission rate of the whole animal and the relative

contribution to the total emission rate by the dissected shells varied between animal

species. M. edulis as the largest species (Tab. 1) showed the highest N2O emission rate

per individual and its shell biofilm contributed on average 94% to the total emission rate.

For L. littorea and H. reticulata, N2O emission from the dissected shell contributed on

average 18% and 32%, respectively, to the total N2O emission rate. In the ZnCl2-treated

incubations of dissected shells, N2O production was not observed (data not shown).

Mytilus edulis

N2O

(nm

ol in

d-1

h-1

)

0,0

0,5

1,0

1,5

2,0

2,5

3,0

Hinia reticulataLittorina littorea

whole animal

dissected shell

Figure 1: N2O emission rates of whole animals and dissected shells of M. edulis, L. littorea andH. reticulata. Freshly collected animals and their dissected shells were incubated in gas-tight vials under oxic conditions with NH4

+ and NO3− amended artificial seawater. The N2O emission

rate was derived from the accumulation of N2O in the incubation vial followed over a period of 4 h. Mean rates ± SD are shown (n = 4-8).

98


15N-stable isotope experiments with dissected shells

15NH4+, 15NO3

− and 15NO2− were all used as precursors for N2O production, revealing

nitrification, denitrification, and denitrification of nitrite as potential sources of N2O in

shell biofilms (Fig. 2, Tab. 3). In the 15NH4+ incubations (targeting nitrification), 46N2O

was produced in biofilms of all three species at average rates ranging from 0.048 to

0.072 nmol g−1 h−1 (Fig. 2a-c, Tab. 3). Nitrification contributed thus 43%, 39% and 47%

to the total 46N2O production (i.e., nitrification plus denitrification) in shell biofilms of

M. edulis, L. littorea and H. reticulata, respectively. 45N2O production in the shell

biofilms of M. edulis started after an initial lag phase of 3 h with a rate 1.5 times higher

than that for 46N2O. In H. reticulata shell biofilms, the 45N2O production rate was 12

times higher than that for 46N2O, whereas in the shell biofilm of L. littorea no 45N2O

production was observed.

In the 15NO3− incubations (targeting denitrification), all three species produced 46N2O at

average rates ranging from 0.075 to 0.080 nmol g−1 h−1 (Fig. 2d-f, Tab. 3).

Denitrification contributed 57%, 61% and 53% to the total 46N2O production

(i.e., nitrification plus denitrification) in shell biofilms of M. edulis, L. littorea and

H. reticulata, respectively. Production of 45N2O was detected in M. edulis shells at a

low rate after a lag phase of 3 h and in H. reticulata shells at a rate 4 times higher than

the 46N2O production rate, indicating the presence or production of 14NO3− and/or

14NO2− in the shell biofilm.

In the 15NO2− incubations (targeting denitrification of nitrite), both 46N2O and 45N2O

production rates of all three species were increased by a factor of 2 to 11 compared to

the respective rates in the 15NO3− and 15NH4

+ incubations (Fig. 2g-i, Tab. 3). In all 15N

incubations, H. reticulata showed the highest N2O production rates of the three species

and 45N2O production rates always exceeded 46N2O production rates, whereas the other

two species produced more 46N2O than 45N2O.

99


0

2

4

6

8

10

15NH4+

15NO3-

15NO2-

c

Mytilus edulis Littorina littorea Hinia reticulata

0,0

0,1

0,2

0,3

0,4

0,5

45N2Ob

N2O

(n

mo

l g

-1)

0,0

0,1

0,2

0,3

0,4

0,5

46N2O

a

N2O

(n

mo

l g

-1)

0,0

0,1

0,2

0,3

0,4

0,5d

0,0

0,1

0,2

0,3

0,4

0,5e

0

2

4

6

8

10f

Time (h)

0 1 2 3 4 5 6

N2O

(n

mo

l g

-1)

0

2

4

6

8

10g

Time (h)

0 1 2 3 4 5 6

0

2

4

6

8

10h

Time (h)

0 1 2 3 4 5 6

0

2

4

6

8

10i

Figure 2: Production of 45N2O (white circles) and 46N2O (black circles) in shell biofilms of M. edulis, L. littorea and H. reticulata. Dissected shells were incubated in 15NH4

+, 15NO3− and

15NO2− amended seawater and sampled over a period of 6 h. Averages of three replicate time

series ± SD are shown. Note different scales on y-axis between panels on white and grey background.

Table 3: 45N2O and 46N2O production rates of dissected shells from incubations with 15NH4+,

15NO3− or 15NO2

−. Mean rates (±SD) are shown, n = 3, n.d. = not detectable.

Rate (nmol g−1 h−1) Mytilus edulis Littorina littorea Hinia reticulata

15NH4+

45N2O 0.085 (0.043) n.d. 0.879 (0.216) 46N2O 0.057 (0.020) 0.048 (0.011) 0.072 (0.015)

15NO3−

45N2O 0.031 (0.014) n.d. 0.313 (0.041) 46N2O 0.075 (0.002) 0.076 (0.007) 0.080 (0.034)

15NO2−

45N2O 0.124 (0.002) 0.060 (0.019) 1.426 (0.251) 46N2O 0.282 (0.095) 0.375 (0.140) 0.787 (0.127)

100


N2O

pro

du

ctio

n r

ate

(n

mo

l N

g-1

h-1

)

0

2

4

6

8

10

Mytilus edulis Hinia reticulataLittorina littorea

Po

ten

tia

l n

itri

ficatio

n

or

de

nitri

fica

tio

n r

ate

(n

mo

l N

g-1

h-1

)

0

20

40

60

80

100

Nitrification assay

Denitrification assay

b)

a)

Nitrification assay

Denitrification assay

Nitrous oxide production in shell biofilms in relation to nitrification and denitrification

Shells of all three species produced N2O in both the nitrification and the denitrification

assay (Fig. 3a). N2O yields of denitrification (percentage of N2O produced per NO3−

consumed) were 13.4%, 11.9% and 5.7%, and of nitrification (percentage of N2O

produced per NH4+ consumed) 3.7%, 7.1% and 4.0% for shells of M. edulis, L. littorea

and H. reticulata, respectively (Fig. 3a+b). All potential N2O production rates were in

the range of 0.20 to 1.05 nmol N g−1 h−1, except for the N2O production rate of

H. reticulata in the denitrification assay which was 4.84 nmol N g−1 h−1. The potential

total denitrification rate of H. reticulata shells was also exceptionally high compared to

the potential rates of nitrification and denitrification of the other two species (Fig. 3b).

M. edulis and L. littorea exhibited higher nitrification than denitrification potentials,

whereas the opposite was found for H. reticulata.

Figure 3: a) Potential N2O production rates of dissected shells from M. edulis, L. littorea and H. reticulata in nitrification (black bars) and denitrification (grey bars) assays. b) Potential nitrification rates (NO2

−

production) (black bars) and denitrification rates (N2+N2O production) (grey bars) of dissected shells from the three species. In the nitrification assays, dissected shells were incubated under oxic conditions with 50 μM NH4

+

for 4 h, in the denitrification assays under anoxic conditions with 50 μM NO3

−

for 4 h. Means ± SD are shown (n = 3). All rates are expressed per mol N. Note different scales on y-axis.

101


Mytilus edulis

Dis

tan

ce

fro

m s

he

ll (m

m)

0,0

0,5

1,0

1,5

2,0

2,5

LightDark

Littorina littorea

Dis

tan

ce

fro

m s

he

ll (m

m)

0,0

0,5

1,0

1,5

2,0

2,5

Hinia reticulata

Oxygen (µM)

0 200 400 600 800 1000 1200 1400

Dis

tan

ce

fro

m s

he

ll (m

m)

0,0

0,5

1,0

1,5

2,0

2,5

Oxygen concentration in shell biofilms

Oxygen concentration gradients inside the shell biofilm varied depending on the light

conditions and thickness of the biofilm (0.05 mm to >1 mm) (Fig. 4). At high light

intensity, the oxygen concentration in the shell biofilms corresponded to 100-500% air

saturation, indicating net oxygen production inside the biofilms. In the dark, the oxygen

concentration in the shell biofilms corresponded to 0-63% air-saturation, indicating net

oxygen consumption inside the biofilms.

Figure 4: Vertical profiles of the oxygen concentration in shell biofilms of M. edulis, L. littoreaand H. reticulata under light (white circles) and dark (grey circles) conditions as measured with microsensors. For each species and light condition, 4 representative profiles are shown that demonstrate the heterogeneity of the oxygen concentration at randomly chosen positions in the shell biofilm.

102



N2O production rates of dissected shells of H. reticulata showed treatment-specific

changes during the incubation in sediment-microcosms for 53 days (Fig. 5, black

squares). For animals with a natural shell biofilm at the beginning of the experiment

(A+ microcosm), the N2O production of their dissected shell increased with time. For

animals with the shell biofilm removed before the experiment (A- microcosm), a

smaller, but steady increase in N2O production of their dissected shell was observed. In

contrast, the N2O production rate of biofilm-covered shells from which the animal was

removed before the experiment (S+ microcosm) remained constant until day 33 and

then decreased to the low level of N2O production of shells from which both the biofilm

and the animal were removed before the experiment (S- microcosm). Thus, the presence

of the animal increased the N2O production potential of the shell biofilm over time.

Also the protein content of shell biofilms developed differently in the four sediment-

microcosms during the incubation period (Fig. 5, grey circles). In the A+ microcosm,

the protein content of the natural shell biofilm stayed constant, whereas in the S+

microcosm, it decreased. In the A- and S- microcosms in which the shell biofilms were

removed before the experiment, the protein content increased slightly or remained at a

very low level, respectively. Thus, the presence of the animal sustained the growth of

the shell biofilm, whereas in the absence of the animal, the biomass of an established

biofilm decreased and no significant biofilm formation on clean shell surfaces could be

observed. Furthermore, the dissolved inorganic nitrogen (DIN) concentration in the

water column increased more in sediment-microcosms with animals than in microcosms

with dissected shells only (Fig. 5, white triangles).

The correlation analysis of the complete data set (4 microcosms x 5 sampling days x 4

replicate N2O measurements, n = 80) revealed that the N2O production rate of dissected

shells was significantly positively correlated with the protein content of the shells

(correlation coefficients in Supplementary Table 2). Moreover, the N2O production rate

of dissected shells was significantly positively correlated with the DIN concentration in

the water column of the microcosms. Additionally, the protein content and the DIN

concentration were also positively correlated with each other.

103


N2O

(n

mo

l g

-1h

-1)

Pro

tein

(m

g g

-1 s

he

ll)

0

2

4

6

8

10

12

14

N2O

Protein

DIN

DIN

(µ

mo

l L

-1)

0

50

100

150

200

250

300

350

400

450

Time (d)

0 5 10 15 20 25 30 35 40 45 50 55

N2O

(n

mo

l g

-1h

-1)

Pro

tein

(m

g g

-1 s

he

ll)

0

2

4

6

8

10

12

14

Time (d)

0 5 10 15 20 25 30 35 40 45 50 55

DIN

(µ

mo

l L

-1)

0

50

100

150

200

250

300

350

400

450

(A+) Animal + Biofilm (A-) Animal - Biofilm

(S+) Shell + Biofilm (S-) Shell - Biofilm

Figure 5: Sediment-microcosm incubation of H. reticulata. A+) animals with natural shell biofilm, A-) animals with the shell biofilm removed before the experiment, S+) dissected shells with natural biofilm, and S-) dissected shells with the biofilm removed before the experiment. N2O production of dissected shells (black squares) was measured on day 1, 20, 33, and 53 by measuring N2O accumulation in gas-tight vials over a period of 4 h. Mean rates of four replicate N2O measurements per microcosm ± SD are shown. Protein contents of the shell biofilms are presented as grey circles and the DIN concentration (sum of NH4

+, NO2− and NO3

−) in the water column of the sediment-microcosm are presented as white triangles.

Discussion

Contribution of the shell biofilm to total N2O emission by marine molluscs

The shell biofilm on three marine mollusc species with different life styles and collected

in different habitats contributed significantly (18-94%) to the total N2O emission of the

animals. On average, N2O production in shell biofilms of marine molluscs is thus in the

same order of magnitude as N2O production inside the animal body. This evidence for

104


substantial N2O production in shell biofilms of abundant marine molluscs may explain

why a recent survey of marine invertebrates revealed a general correlation between the

N2O emission rate and the presence of a microbial biofilm on exoskeleton and shell

surfaces (Heisterkamp et al., 2010). These results complement earlier studies on N2O

emission from terrestrial and aquatic invertebrates that ascribed N2O production

exclusively to microbial denitrification activity in the anoxic gut (Drake and Horn, 2007;

Stief et al., 2009). The extreme example here is the blue mussel M. edulis, in which N2O

production originated almost exclusively from the shell biofilm. Apparently, N2O

production by gut denitrification is negligible in this species. This is surprising, since M.

edulis is a very efficient filter-feeder that ingests large amounts of bacteria (McHenery

and Birkbeck, 1985) and is thus likely to promote high N2O production in its gut (Stief

et al., 2009). However, the high lysozyme activity in the gut of M. edulis (Birkbeck and

McHenery, 1982) might digest most of the ingested bacteria and thereby inhibit

denitrification and concomitant N2O production in the gut. It can be speculated that also

in other aquatic species than M. edulis that are listed in Stief et al. 2009 N2O emission

might originate at least partly from shell biofilms. For the freshwater mussel Dreissena

polymorpha, it has recently been shown that the shell biofilm contributes about 25% to

the total N2O emission of the animal (Svenningsen et al., 2012).��

N2O emission from L. littorea and H. reticulata derived only partly from N2O

production in the shell biofilm, while the majority of N2O was produced in other parts

of the animal body. Besides the gut, the gills especially may be sites of N2O production,

as they exhibit nitrification activity in various mollusc species (Welsh and Castadelli,

2004).

N2O-producing pathways in shell biofilms

The stable isotope experiments revealed that nitrification and denitrification produce

N2O in shell biofilms of all three mollusc species. Incubations with the tracers 15NH4+,

15NO3− and 15NO2

− were designed to specifically ascribe the production of the double-

labelled 46N2O to nitrification, denitrification of nitrate and denitrification of nitrite,

respectively. Based on the 46N2O production rates in the 15NH4+ and 15NO3

− treatments,

nitrification contributed on average 43% and denitrification 57% to the total 46N2O

production. Both processes are thus almost equally important for N2O production in

105


shell biofilms of marine molluscs. Consequently, N2O emission from marine

invertebrates can originate from at least two microbial processes in two different body

compartments: denitrification in the gut of the animal (Heisterkamp et al., 2010, Stief et

al., 2009), and from denitrification as well as nitrification in the shell biofilms. This

means that besides nitrate and nitrite, ammonium also serves as a precursor and

consequently as regulating factor for animal-associated N2O production.

In contrast to the 46N2O production rates, which were similar for all three species,

biofilms differed in their 45N2O production rates. In the 15NH4+ treatment, 45N2O

production rates show that only in shell biofilms of H. reticulata nitrification and

denitrification were tightly coupled and/or that nitrifier denitrification occurred at a

significant rate. In shell biofilms of M. edulis, coupled nitrification-denitrification and

nitrifier denitrification were apparently limited by the ammonium oxidation step, as

indicated by the lag-phase in 45N2O production, whereas in shell biofilm of L. littorea

no mixing of the 14N and 15N pools took place. The cell-to-cell connectivity and mixing

of products with the ambient pools seem thus to differ between biofilms on different

species. Shell biofilms of H. reticulata produced also high amounts of 45N2O in the 15NO3

− treatment, although no additional 14N pool was added and the background 14N concentration in the artificial seawater was about 5 μM DI14N. Provided that

random isotope pairing occurred according to the 14N to 15N ratio of 1:10 ([5×14N]2 +

2[5×14N][50×15N] + [50×15N]2), the 45N2O production rate should be 20% of the 46N2O

production rate. The far higher 45N2O production rate in H. reticulata shell biofilms

suggests that more than 5 μM DI14N was present. The additional 14N probably

originated from the biofilm itself, either stored in cells or bound to the extracellular

matrix of the biofilm or produced by remineralization during the incubation period.

Apparently, this internal 14N pool is more easily available for the cells than the external 15N pool, thereby enhancing 45N2O production disproportionately despite the much

larger 15NO3− pool in the incubation medium.

The 15NO2− incubations revealed that nitrite strongly increased the rates of 46N2O and

45N2O production of all three species compared to the respective rates in the incubations

with equimolar concentrations of 15NH4+ and 15NO3

−. Furthermore, the low 45N2O/46N2O ratios, despite the 14N to 15N ratio of 10:1, show that the nitrite pool is

more readily used than the nitrate pool in shell biofilms of all three species. Chemical

106


conversion of NO2− to N2O can be ruled out as an explanation for the higher N2O

production rates in the presence of nitrite, since the negative controls with killed

dissected shells incubated in 50 μM and 500 μM NO2− did not result in a significant

accumulation of N2O (Supplementary Fig. 1). Instead, nitrite might have enhanced

biological N2O production due to increased nitrifier denitrification activity. Ammonia-

oxidizing bacteria and complex biofilms produce high amounts of N2O when

denitrifying nitrite at low oxygen concentration or high nitrite concentration (Wrage,

2001; Beaumont et al., 2004; Schreiber et al., 2009). Likewise, nitrifiers in shell

biofilms might have been stimulated to reduce nitrite to N2O by low oxygen

concentration or elevated nitrite concentration in the biofilm. A nitrite concentration of

50 μM is unlikely to be toxic to bacteria (Stein and Arp, 1998; Tan et al., 2008), but

bacteria may increase nitrite reduction rates already at low nitrite concentrations to

avoid that toxic levels are being reached.

The total N2O production rate is a function of the overall process rate of nitrification

and denitrification and the N2O yield of these processes (percentage of N2O production

per turn-over of substrate). The N2O yields from nitrification and denitrification in shell

biofilms were relatively high (i.e., 3.7-13.4%) compared to N2O yields from water

column nitrification (de Wilde and de Bie, 2000), sedimentary denitrification in

eutrophic estuaries (Dong et al., 2006), marine ammonia-oxidizing bacteria (AOB)

(Frame and Casciotti, 2010), and marine ammonia-oxidizing archaea (AOA) cultures

(Santoro et al., 2011). They were, however, in the same range as N2O yields of

denitrification from rocky biofilms in an intertidal area (Magalhaes et al., 2005). In

contrast to the stable isotope experiments in which dissected shells were incubated at

initially air-saturated conditions, the potential rates of nitrification and denitrification

and their N2O yields were measured under completely oxic (continuously aerated) or

anoxic atmosphere (sealed). These conditions were thus optimal for nitrification or

denitrification, leading probably to higher process rates and consequently higher N2O

production rates than in the stable isotope incubations. However, as these potential rates

were measured under completely oxic or anoxic conditions, the N2O yields presented

here might be underestimates, since N2O yields of nitrification and denitrification are

generally highest under low oxygen concentrations (Goreau et al., 1980; Jørgensen et al.,

1984; Bonin and Raymond, 1990). Conversely, at low oxygen and substrate

concentrations, the rates of nitrification and denitrification are likely to slow down

107


(Jørgensen et al., 1984; Codispoti et al., 2005) and may thus counteract the effect of an

increased N2O yield, leading to an only moderate increase in N2O production under

conditions suboptimal for either process.

The oxygen distribution in the shell biofilm was very heterogeneous, varying with light

intensity and thickness of the biofilm. At low light intensities, nitrification and

denitrification probably co-occur in the shell biofilm, with nitrification taking place in

the (fully) oxic surface layer and denitrification in the hypoxic or anoxic bottom layer of

the biofilm. At high light intensities, the biofilm is completely oxic and denitrification

only takes place if bacteria capable of aerobic denitrification are present. Several

bacterial strains are able to denitrify at or above air saturation (Patureau et al., 2000;

Zehr and Ward, 2002; Hayatsu et al., 2008) and high rates of aerobic denitrification

were measured in permeable intertidal sediments which are very dynamic environments

with changing oxygen concentrations (Gao et al., 2010). The oxygen distribution at high

light intensities allows nitrification to occur throughout the complete biofilm, but the

rate of nitrification might be reduced as nitrifiers are known to be inhibited by high light

intensities (Horrigan and Springer, 1990).

The biofilms on living animals are influenced by the respiration, feeding and migration

behaviour of the animal, which expose the shell biofilm to changing environmental

conditions, thereby leading to frequent changes in the oxygen concentration inside the

shell biofilm. Fluctuations in oxygen concentration result in high transient N2O

production by AOB and denitrifiers in pure cultures and microbial biofilms (Kester et

al., 1997; Bergaust et al., 2008; Schreiber et al., 2009). Similarly, shell biofilms are

presumably sites of high N2O production under in situ conditions.


The presence of the animal enhanced N2O production in the shell biofilm by stimulating

biofilm growth and providing a nutrient-enriched environment. The protein content of

the dissected shells (used as a proxy for biofilm biomass) and the water column DIN

concentration were increased in microcosms with animals compared to microcosms

with shells only and were both significantly positively correlated with the N2O

production rate of the shell biofilms. The animals increased the water column DIN

108


concentration probably due to their feeding activity and high ammonium excretion rate

as well as stimulation of DIN release from the sediment by bioturbation (Aller et al.,

2001). Furthermore, the ammonium excretion rates of H. reticulata as well as of

M. edulis and L. littorea (Table 1) are high enough to support the measured total N2O

production in shell biofilms assuming that nitrification and denitrification are tightly

coupled. In this case, animal-associated N2O production can be sustained by the

excretion of the animal alone and is thus independent from an ambient DIN source.

A stimulating effect of increased DIN on N2O production was also reported for rocky

biofilms (Magalhaes et al., 2005) and marine sediments (Seitzinger and Kroeze, 1998),

and was attributed to increased denitrification rates and/or increased N2O yields under

elevated nutrient concentration. However, only a minor effect of the DIN concentration

on the N2O production rate was observed during short-term incubations (4 h) of

dissected shells (Supplementary Information Table 1). Thus, only the long-term

exposure to different nutrient concentrations (days to weeks) seems to affect N2O

production in shell biofilms, probably by influencing the microbial abundance and

community composition. In intertidal biofilms, elevated nutrient concentrations in the

water column increase biofilm density and change the composition of its bacterial

community (Chiu et al., 2008). The significant positive correlation between the DIN

concentration and the protein content of the shell further substantiates that the elevated

DIN concentration due to the presence of the animal determines growth and probably

also the microbial composition of the biofilm.

Conclusion

N2O production in shell biofilms of marine molluscs originates from both nitrification

and denitrification and contributes significantly (18-94%) to the total N2O emission

from the different species of marine molluscs tested in this study. Particularly high N2O

production occurs during denitrification of nitrite. Animal-associated N2O production

can thus be fuelled by ammonium, nitrate, and nitrite. The animal stimulates microbial

growth on its shell surface and provides a special micro-environment that is

characterized by high nutrient availability and dynamic changes of oxygen

109


concentration. These conditions favour N2O production by nitrification and

denitrification in shell biofilms compared to microbial biofilms not directly affected by

animals. Many marine taxa possess a complex microbial biofilm on their external

surfaces (Wahl, 1989) and might provide similar habitats in which N2O production is

stimulated. N2O emission from invertebrates is thus likely to be widespread in marine

environments and of particular importance in areas with high animal abundance, such as

natural beds or longline farms of the blue mussel M. edulis.

Acknowledgements

We are grateful to Anna Behrendt for assistance in the laboratory and to Lars Peter

Nielsen for advice and support. This work was financially supported by the DFG grant

STI202/6-1 awarded to P.S. and by the Max Planck Society.

110


Supplementary Information

Dissected shells of H. reticulata were incubated for 4 h at different DIN concentrations

to test for short-term effects of the DIN concentration on N2O production rates in shell

biofilms. On day 53 of the sediment-microcosm experiment, dissected shells of all four

treatments were incubated in filtered seawater from the respective microcosm and N2O

production was followed over 4 h by taking headspace samples. Additionally, dissected

shells from the treatments A+ and A- were incubated with filtered seawater from the S-

microcosm which contained a DIN concentration of 136 μM (1 μM NH4+, 1 μM NO2

−,

134 μM NO3−), while the shells of the treatments S+ and S- were incubated with filtered

seawater from the A+ microcosm that contained 456 μM DIN (8 μM NH4+, 22 μM

NO2−, 426 μM NO3

−). This reciprocal 4h-incubation of A+ and A- shells in S- seawater

and of S+ and S- shells in A+ seawater did not result in significant differences in the

N2O production rates of A+, A- and S+ shells (Supplementary Table 1). Only the N2O

production rate of S- shells was significantly higher when incubated with A+ seawater

instead of S- seawater. The DIN concentration had thus an only minor effect on the N2O

production rate during short-term incubation of 4 h.

Supplementary Table 1: t-test comparison of the N2O production rates in the reciprocal 4h-incubation of A+ and A- shells in seawater from the respective microcosm and in S- seawater, and of S+ and S- shells in seawater from the respective microcosm and in A+ seawater.

P t Df

A+ 0.228 -1.44 3.75

A- 0.925 0.101 3.94

S+ 0.850 0.20 4.52

S- 0.044 -2.78 4.52

111


Supplementary Table 2: Correlation coefficients between N2O production rate of dissected shells (nmol g−1 h−1), protein content of dissected shells (mg g−1) and DIN concentration in the water column of the microcosms (μM). n = 80 (4 microcosms x 5 sampling days x 4 replicate N2O measurements).

Linear correlation

(Pearson coefficient)

Non-parametric correlation

(Spearman coefficient)

N2O N2O

Protein 0.478 (p < 0.001) 0.684 (p < 0.001)

DIN 0.639 (p < 0.001) 0.596 (p < 0.001)

Protein Protein

DIN 0.381 (p < 0.001) 0.335 (p = 0.002)

Dissected shell + 50 µM NO2

-

Dissected shell + 500 µM NO2

-

Autoclaved artificial seawater + 500 µM NO2

-

Autoclaved artificial seawater + 50 µM NO2

-

N2O (nmol g

-1 h

-1)

0,0 0,5 1,0 1,5 2,0 6,0 7,0 8,0 9,0

Autoclaved dissected shell + 50 µM NO2

-

Autoclaved dissected shell + 500 µM NO2

-

Supplementary Figure 1: Negative controls were performed to test for chemical conversion of NO2

− to N2O. First, autoclaved artificial seawater amended with either 50 or 500 μM NO2− was

incubated for 6 h and analyzed for N2O production by gas chromatography. Second, autoclaved dissected shells were added to the autoclaved artificial seawater amended with either 50 or 500 μM NO2

− and then analyzed for N2O production. In none of the negative controls a significant increase in N2O could be detected during the incubation period of 6 h. Thus, chemical conversion of NO2

− to N2O can be ruled out as an explanation for the very high N2O production by live shell biofilms.

112


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��

Chapter 4

Cover photograph (Copyright © 2012, American Society for Microbiology. All Rights Reserved.): Close-up of a zebra mussel (Dreissena polymorpha) reef. This species is invasive in North American and European freshwater systems and can form reefs of more than 100,000 individuals per square meter. Nitrification in shell biofilms and denitrification in the mussel's gut may dramatically increase benthic N2O emissions.

��

117

�

�118

��

Shell biofilm nitrification and gut denitrification contribute to emission of nitrous oxide

by the invasive freshwater mussel Dreissenapolymorpha (Zebra Mussel)

Nanna B. Svenningsen1, Ines M. Heisterkamp2, Maria Sigby-Clausen1,

Lone H. Larsen1, Lars Peter Nielsen1, Peter Stief2, and Andreas Schramm1





Applied and Environmental Microbiology 78(12): 4505-4509, 2012

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Chapter 4 N2O emission from Dreissena polymorpha�

Abstract

Nitrification in shell biofilms and denitrification in the gut of the animal accounted for

N2O emission by Dreissena polymorpha (Bivalvia), as shown by gas chromatography

and gene expression analysis. The mussel’s ammonium excretion was sufficient to

sustain N2O production and thus potentially uncouples invertebrate N2O production

from environmental N concentrations.

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Introduction

Nitrous oxide (N2O) is a powerful greenhouse gas that contributes to stratospheric

ozone destruction (3, 4). In natural systems, the production of N2O is primarily

associated with the turnover of inorganic nitrogen compounds by nitrifying and

denitrifying microorganisms, often in oxic/anoxic transition zones in soil and sediment

(34). Nitrifiers (both ammonia-oxidizing bacteria and archaea) produce N2O as a by-

product of ammonia oxidation (6, 28), especially under oxygen limitation, while for

denitrifiers N2O is an intermediate in anaerobic respiration (40). Besides soils and

aquatic systems, also invertebrates are sites of a globally significant N2O production,

first discovered for earthworms (15, 19), and subsequently for diverse freshwater and

marine invertebrates (11, 36). This animal-associated N2O production has been

attributed to incomplete denitrification by ingested microorganisms in the anoxic

invertebrate gut (13, 14, 35). In addition, biofilms covering shells and exoskeletons of

marine invertebrates have been identified as sites of N2O emission (11). Their relative

contribution to animal-associated N2O production, the pathways involved, and their

distribution among marine and freshwater invertebrates are still unknown. The objective

of the present study was therefore to quantify the biofilm-derived N2O production and

its mechanism(s) using the N2O-emitting (36) freshwater bivalve Dreissena polymorpha

(zebra mussel) as model organism. This species is considered invasive in North

America and Europe, and can occur at extremely high abundance. Local populations in

the Gudenå river system (Denmark) occasionally form large reefs at the sediment

surface with more than 100,000 individuals per m2 (1).

Methods, Results, and Discussion

Site of N2O production in D. polymorpha

Mussels were sampled in April 2010 in the river Remstrup, which is part of the Gudenå

system. Living animals or shells dissected from living animals were pooled in sets of

7-15 individuals for replicate incubations (n = 5-6) at 21�C in gas-tight bags (10) filled

with air-saturated artificial freshwater (33) containing NH4+ and NO3

- (50 μM each) and

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a headspace of atmospheric air. Shells incubated with 50 % ZnCl2 to kill biological

activity served as negative controls. N2O emission rates were determined from linear

increase of N2O concentrations in 3–h incubations as previously described (36). In

short, water samples were hourly withdrawn from the bags, transferred to N2-flushed,

ZnCl2-containing exetainers, and N2O was measured by gas chromatography (36). Bags

were still oxic (>50 %) after the 3-h incubation as confirmed with an O2 microelectrode

(26).

N2O emission was approximately linear over time both in incubations of whole mussels

and biofilm-covered dissected shells; for whole mussels, the rates were similar to

D. polymorpha collected in August 2006 in the river Rhine (36). The shell biofilm

contributed approximately 25% to the total N2O emission from D. polymorpha

specimens (Fig. 1). N2O production was an exclusively biological process, indicated by

the linearity of the emissions and confirmed by the absence of N2O emissions in the

killed control.

Figure 1: N2O emission from living animals or shells dissected from living animals incubated in artificial freshwater with (+ ATU) or without inhibition of NH3 oxidation by ATU (Ctrl). Error bars represent standard deviations (SD) of the mean (n=5-6, each replicate consists of 7-15 animals or shells). Different lowercase letters indicate significant differences between treatments (p< 0.05, t-test)

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Pathways of N2O production

Additional whole animals and dissected shells were incubated with allylthiourea (ATU,

100 μM) to inhibit NH3 oxidation (8). N2O emission from ATU-incubated shells was

almost completely eliminated, pointing to nitrification as the dominant N2O-producing

pathway in the shell biofilm of D. polymorpha (Fig. 1). In contrast, N2O emission from

the animal itself was not reduced by ATU, which indicates that denitrification was

responsible for N2O production inside the animal, in agreement with gut-associated N2O

production via denitrification in other freshwater invertebrates (36).

These results were supported by the detection of transcripts for bacterial ammonia

monooxygenase (amoA), the key enzyme of ammonia oxidation, and for nitrite

reductase (nirK and nirS), a key enzyme of denitrification. RNA was extracted from

dissected, whole guts and from biofilm material (sampled in June 2010) with the

FastRNA® Pro Soil-Direct Kit (MP Biomedicals), and DNase-treated (Ambion) for

30 min to remove DNA, as confirmed by (lack of) 16S rRNA gene-specific PCR

amplification. Reverse transcription PCR (RT-PCR, 35 cycles) was performed with the

OneStep RT-PCR Kit (Qiagen). Published protocols and primers specific for bacterial

amoA, amoA1F-amoAR-TC (24, 27) and for nirK and nirS, F1aCu-R3Cu (9) and

Cd3aF-R3cd (21, 38), respectively, were used. Bacterial amoA mRNA was only

detected in biofilm samples, while mRNA of nirK and nirS were only detected in gut

samples (Table 1). Since archaeal amoA genes were never detected by PCR (12) in any

of the samples (see below), detection of archaeal amoA transcripts was not attempted.

Additional animals were collected in December 2010 and analyzed by reverse

transcription quantitative PCR (RT-qPCR). Mussels were incubated for 4 hours at

similar conditions as during N2O rate measurements. Then total nucleic acids were

extracted in triplicate by a phenol-chloroform protocol (7, 25), and one aliquot of the

nucleic acid extract was DNase-treated as described above. cDNA synthesis with the

Omniscript Reverse Transcription kit (Qiagen) was primed by random hexamers, and

cDNA copy numbers of bacterial amoA, nirK and nirS were quantified in a LightCycler

480 (Roche) as described previously (12). Annealing temperatures were adjusted to

55�C for nirS and to 57�C for bacterial amoA and nirK; detection limit (10-13 cDNA

copies) was defined as 3x the standard deviation of the non-template control, while the

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limit of quantification was defined by the lower limit of the linear range of the standard

curves (85-100 cDNA copies).

Copy numbers of all cDNAs were low (always below the limit of quantification, for

nirS always below the detection limit), but confirmed the results of the qualitative

RT-PCR assay for bacterial amoA and nirK: bacterial amoA cDNA was only detected in

biofilm samples, while nirK cDNA was only detected in gut samples (Table 1).

Table 1: Expression of genes encoding ammonia monooxygenase (amoA) and nitrite reductase (nirK and nirS) in animals collected in June and December 2010. Expression ofa:

amoA nirK nirS

Material June

December (cDNA copies per mg wet-weight) June

December (cDNA copies per mg wet-weight) June

December cDNA copies per mg wet-weight)

Gut � � + 205-1585b + < 240c

Shell biofilm

+ 200-2000b � � � �

a �, not detected by RT-PCR or RT-qPCR; +, detected by RT-PCR. b Above limit of detection but below limit of quantification for RT-qPCR. c Below limit of detection for RT-qPCR but detected and cloned after RT-PCR.

To test for the metabolic potential of the biofilm and gut microbial community, gene

copy numbers of bacterial amoA, nirK and nirS were quantified in the nucleic acid

extracts from December 2010. Amplification of archaeal amoA (12) was attempted

several times, but the result was always negative, indicating that archaeal ammonia

oxidizers were not relevant in these samples. qPCR were performed as described above,

and functional gene copy numbers were normalized against 16S rRNA gene copy

numbers amplified with primer pair 341F-907R (22, 23), with annealing at 57 �C.

16S rRNA gene copy numbers (per mg wet weight) were 4.83 × 106 ± 6.2 × 105 in the

gut and 3.48 × 108 ± 1.7 × 107 in the shell samples. Copy numbers of all functional

genes were above the limit of quantification. Relative abundance ± SD was low in gut

samples, i.e., 1.6 × 10�3 ± 1.5 × 10�3 for bacterial amoA, 2.7 × 10�1 ± 6.5 × 10�2 for

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nirK, and 2.5 × 10�1 ± 3.8 × 10�2 for nirS. Biofilm samples showed higher relative

abundances, i.e., 2.0 × 10�2 ± 2.4× 10�3 for bacterial amoA, 1.6 × 101 ± 9.6 × 10�1 for

nirK, and 1.6 × 100 ± 1.2 × 10�1 for nirS. These data indicate a potential for ammonia

oxidation and denitrification in both gut and biofilm, if environmental conditions allow.

Expression of bacterial amoA, nirK and nirS is affected by a variety of environmental

factors, including O2 partial pressure and availability of N-substrates (29, 40). Inside the

mussel gut, O2 will most likely be depleted (35). In accordance with the data presented

here, denitrification will therefore be induced, and ammonia oxidation repressed, when

denitrifiers and ammonia oxidizers, respectively, enter the gut. Mussel biofilms

analyzed in this study, on the other hand, were relatively thin and presumably fully oxic,

as indicated by preliminary O2-microsensor measurements (data not shown). N2O is

therefore mainly produced by nitrification, while denitrification is repressed. However,

high nir gene abundance indicates that denitrification may contribute to N2O

production, if anoxic microsites develop within the biofilm (30).

Diversity of expressed amoA, nirK, and nirS

To assess the diversity of the active ammonia oxidizers and denitrifiers, clone libraries

were constructed from cDNA of bacterial amoA (biofilm samples), and nirK/nirS (gut

samples) of animals collected in June and December 2010. RT-PCR products were

cloned using the pGEM-T cloning kit (Promega), and approx. 30 randomly picked

clones per sample and gene were sequenced (GATC Biotech; Macrogene); the cDNA

clone sequences were deposited in Genbank under accession numbers JF820296-

JF820311. Sequences were aligned by the integrated aligner tool in the ARB software

(18) together with sequences of their closest relatives found by nucleotide BLAST,

translated into amino acid sequences and used for phylogenetic tree construction in

ARB using neighbor joining and maximum likelihood analysis with 1000 bootstraps

replications. Both methods resulted in identical tree topologies.

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Figure 2: Neighbor joining tree of amino acid sequences deduced from cDNA obtained in June 2010 (red) or December 2010 (green). Sequences of bacterial amoA (a) are from shell biofilms, while sequences of nirK (b) and nirS (c) are from the gut of D. polymorpha. The number of identical sequences (at least 97 % nucleotide identity) is shown in brackets. Scale bar, 10% amino acid sequence divergence. Node symbols indicate bootstrap support by maximum likelihood analysis: closed circles, >75%; open circles, >50%.

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Sequences of expressed bacterial amoA were in June 2010 affiliated with the

Nitrosomonas europaea- and N. oligotropha lineages, and a lineage without cultured

relatives, while in December 2010 they were affiliated with the Nitrosospira- and

N. oligotropha-lineage (Fig. 2a). Since both clone libraries were well covered (Good’s

coverage >98% based on a 97% nucleotide similarity threshold), the most probable

explanation is differential activity of ammonia oxidizers at the different sampling times,

possibly related to their differing substrate affinities (16).

In contrast, diversity of expressed nirK and nirS was very low, and sequences retrieved

from animals collected in June and December were highly similar or identical. NirK

affiliated with Dechloromonas aromatica (87% DNA sequence similarity), while nirS

were only distantly related (70% DNA sequence similarity) to various

Alphaproteobacteria, e.g. Rhodopseudomonas palustris or Rhodobacter spheroides

(Fig. 2b, c). This limited diversity of active denitrifiers in the gut may be explained by

the fact that mussels are capable of feeding on a diet of bacteria due to high lysozyme

content in their digestive organs (20, 32). Consequently, only a minor part of the

ingested denitrifiers may survive and induce their denitrification genes during the gut

passage in D. polymorpha.

Ammonium excretion by D. polymorpha

The availability of NH3 (as substrate for ammonia oxidation) is usually low in natural

freshwater systems but can be high in environments infested with D. polymorpha

(5, 17). Ammonium excretion rates of D. polymorpha were measured by incubating

groups of 1-8 living mussels (n = 6) in artificial freshwater without amendment of any

N-sources. NH4+ concentrations were quantified spectrophotometrically (2) every half

hour for a total of three hours. The average excretion rate ± SD was 0.128 ± 0.063 μmol

NH4+ individual�1 h�1, which is >1000 times the N needed to explain the N2O

production by nitrification in shell biofilms. Therefore, a significant part of the mussels’

N2O emission is sustained by the animals’ N excretion.

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Environmental implications The results presented here are important on three accounts. First, they provide

quantitative data for the contribution of shell biofilms to the overall N2O emission by a

benthic freshwater invertebrate, hence extending earlier qualitative observations on

marine invertebrates (11). Second, with a substantial part of N2O produced via

nitrification, which can be entirely fuelled by the mussels’ own ammonia excretion, the

data suggest that invertebrate-associated N2O emissions can be decoupled from

environmental nitrate concentrations, one of the main drivers of gut denitrification (19,

37, 38). In addition, biofilm nitrification may not only directly produce N2O but may

also provide nitrate for denitrification-derived N2O production inside the mussel.

The data also show a considerable potential of the invasive D. polymorpha to contribute

to overall N2O emissions from zebra mussel-infested ecosystems. Maximum densities

of up to 100,000 individuals per m2 in the river Gudenå and a potential emission rate of

144 pmol N2O ind�1 h�1 amount to an emission potential of 28 μmol N2O-N m�2 h�1 for

D. polymorpha, or up to 400 times the areal N2O fluxes reported for (non-infested)

freshwater environments (31). Finally, shell biofilms, ammonium excretion, and

coupled nitrification-denitrification are likely to combine also for other freshwater and

marine invertebrates into significant N2O emission potentials (Heisterkamp et al.,

unpublished data). It should however be noted that for assessing their true

environmental impact, in situ studies will be necessary, combining activity

measurements and molecular analyses throughout the seasonal cycle.

Nucleotide sequence accession numbers

cDNA clone sequences obtained in this study have been deposited in GenBank under

accession no. JF820296 to JF820311.

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Acknowledgments

We thank Preben G. Sørensen and Britta Poulsen for expert help in the laboratory. This

study was supported by the Danish Research Council and the German Science

Foundation (grant STI202/6-1 to P.S.).

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Chapter 5

The Pacific White Shrimp Litopenaeus vannamei is a globally important aquacultured species that emits nitrous oxide at a high rate due to incomplete denitrification of ingested bacteria in its anoxic gut.

133

Incomplete denitrification in the gut of the aquacultured shrimp Litopenaeus vannamei

as source of nitrous oxide

Ines M. Heisterkamp1, Andreas Schramm2, Dirk de Beer1, Peter Stief1

1Microsensor Research Group, Max Planck Institute for Marine Microbiology,

Celsiusstraße 1, 28359 Bremen, Germany.


Ny Munkegade 114, 8000 Aarhus C, Denmark

Preliminary manuscript

135

Chapter 5 N2O production in aquacultured shrimp

Abstract

The Pacific White Shrimp Litopenaeus vannamei is a globally important aquaculture

species that is reared at high animal densities, temperatures, and nutrient concentrations.

This species was found to emit the greenhouse gas nitrous oxide (N2O) at the highest

rate recorded so far for any marine invertebrate species. Under in situ conditions of a

recirculating aquaculture farm in Northern Germany (i.e. 28°C and 10 mmol L�1 nitrate),

L. vannamei emitted on average 4.33 nmol N2O per individual and hour. Nitrous oxide

emission of the shrimp derived primarily from N2O production by ingested denitrifying

bacteria in the animal’s gut. The mean N2O yield of gut denitrification (i.e. the fraction

of N2O produced per total nitrogen gas produced) was as high as 36.5%. This high N2O

yield was most probably caused by delayed induction of the N2O reductase after

ingestion of denitrifying bacteria from the oxic water column into the anoxic gut as

demonstrated by oxic-anoxic shift experiments. Inhibition of the N2O reductase by low

oxygen concentrations or pH values was ruled out by microsensor measurements which

revealed that the gut is completely anoxic and has a slightly alkaline milieu. The short

gut passage time of only 1 h apparently prohibits the ingested denitrifiers from

establishing complete denitrification to dinitrogen. In fact, the shrimp guts represent

abundant microsites of incomplete denitrification that significantly contribute to the

N2O supersaturation of on average 2390% in the rearing tanks. In conclusion, microbial

N2O production directly associated with L. vannamei or other aquaculture species

should be considered as important sources of N2O in animal production.

136


Introduction

Aquaculture systems are usually characterized by high loads of nutrients, especially

nitrogen, and accordingly are sites of intense nitrogen turnover, including the microbial

processes of nitrification and denitrification (Crab et al. 2007, Hu et al. 2012). Both

processes can produce nitrous oxide (N2O), which is a potent greenhouse gas and the

major ozone-depleting substance (Forster et al. 2007, Ravishankara et al. 2009).

Aquaculture was therefore recently discussed as an important source of atmospheric

N2O (Williams & Crutzen 2010, Hu et al. 2012). It was estimated that the global N2O

emission from aquaculture currently accounts for 0.09-0.12 Tg N yr�1 and will rise to

0.38-1.01 Tg N yr�1 by 2030 due to the rapidly growing aquaculture industry (William

& Crutzen 2010, Hu et al. 2012). Since capture fisheries has leveled off, aquaculture has

become an important alternative for production of food fish, including finfishes,

crustaceans, molluscs, and other aquatic animals, and has grown with an average annual

rate of 8.3% since 1970 to a total production of 52.5 million tons in 2008 (FAO 2010).

Crustaceans alone accounted for 5 million tons in 2008 of which 2.3 million tons were

made up by one single species, the Pacific White Shrimp Litopenaeus vannamei

(FAO 2010). The maximal estimated N2O emission from aquaculture in 2030 would

represent 18% and 5.7% of the current estimates of the total aquatic and the global N2O

emissions, respectively (Denman et al. 2007). However, so far direct measurements of

N2O emissions from aquaculture are missing and the current estimates are based on the

overall nitrogen load of aquaculture systems and N2O-emission factors that were

derived from wastewater treatment plants (Williams & Crutzen 2010, Hu et al. 2012). It

is highly uncertain whether these emission factors reflect the true N2O yield of nitrogen

cycling processes in aquaculture farms, since very little is known about the mechanisms

and controlling factors of microbial N2O production in aquaculture systems. The

amount of N2O produced by nitrification and denitrification depends on diverse factors

such as oxygen concentration, pH, temperature, and availability of substrates, which can

vary between different aquaculture systems and between different compartments of a

single aquaculture system (Crab et al. 2007, Hu et al. 2012). Nitrification takes place in

the water of the aerated rearing tanks, whereas denitrification occurs in anoxic niches of

the bead bed or in floating organic particles (Holl et al. 2010). In recirculating

aquaculture systems (RAS), biological treatment of the water is necessary to prevent

137


accumulation of nitrogen compounds such as ammonia and nitrite to toxic levels.

Similar to waste water treatments plants, the aquaculture water is processed in external

biofilters, in which nitrification and denitrification prevail at high rates due to massive

microbial biomass in these filters (van Rijn 1996).

Additional microsites of microbial N2O production that so far have been overlooked in

aquaculture systems are represented by the guts of the reared animals. This is in so

much surprising since many free-living terrestrial, freshwater, and marine invertebrate

species are known as ecologically important N2O-emitters (Karsten & Drake 1997, Stief

et al. 2009, Heisterkamp et al. 2010). For earthworms and insect larvae, it has been

shown that the gut microenvironment is characterized by anoxia and the presence of

nitrate and labile organic carbon sources, which stimulates the denitrification activity of

ingested bacteria (Horn et al. 2003, Stief & Eller 2006). The complete denitrification

pathway involves four reduction steps (NO3� � NO2

� � NO � N2O � N2), each of

which is catalyzed by a specific enzyme (Zumft 1997). The N2O reductase plays a key

role because the presence/absence and the activity level of this enzyme determine

whether denitrification acts as a source or sink of N2O. A striking feature of

denitrification in the gut of invertebrates is the high yield of N2O that makes up 15 to

68% of the total nitrogen gas production (N2O+N2), compared to usually less than 1% in

the sediment and water column of rivers, lakes, and coastal aquatic systems (Seitzinger

1988, Drake & Horn 2007, Stief et al. 2009, Beaulieu et al. 2010). Several

environmental factors promote incomplete denitrification by acting upon the expression

and activity levels of the N2O reductase relative to the other three denitrification

enzymes. In general, the N2O/N2 ratio of denitrification will be increased by hypoxic

conditions, low pH, low temperature, high nitrate concentration, and low C/N ratio due

to a low relative activity of the N2O reductase (Bonin & Raymond 1990, Richardson et

al. 2009, Bergaust et al. 2010, Hu et al. 2012). Furthermore, more N2O will be produced

than consumed, if the denitrifier community comprises many bacterial strains that do

not possess the N2O reductase gene and thus lack the ability to reduce N2O to N2 (Zumft

1997, Gregory et al. 2003). Additionally, the delayed induction of the N2O reductase

after sudden shifts from oxic to anoxic conditions may cause (transiently) high N2O

yields (Baumann et al. 1996, Kester et al. 1997). This scenario was hypothesized to

occur during the feeding process of invertebrates, which abruptly transfers facultative

138


denitrifying bacteria from the ambient oxic environment into the anoxic gut (Drake et al.

2006, Stief et al. 2009).

To date, measurements of in situ N2O emission rates from aquacultured invertebrates

and detailed investigations on the factors controlling the N2O yield of denitrification in

the invertebrate guts are missing. This study therefore aimed at elucidating the in situ

N2O emission rate of the aquacultured Pacific White Shrimp Litopenaeus vannamei

(also Penaeus vannamei, Boone 1931), and the major factors that might regulate the

N2O yield of gut denitrification in this species. The penaeid shrimp L. vannamei is the

most important crustacean species in aquaculture worldwide and is reared in intensive

aquaculture systems at high stocking densities, temperatures, and nutrient

concentrations (Cuzon et al. 2004, Browdy & Jory 2009, FAO 2010).


Nitrous oxide emission rates from whole animals and dissected guts

L. vannamei specimens were obtained from a RAS in Northern Germany, in which they

were reared at a temperature of 29.5 ± 0.5°C, pH of 8.08 ± 0.14, salinity of 19 ± 2,

dissolved organic carbon concentration of 20.6 ± 7.0 mg L�1, and a nitrate concentration

of 9.13 ± 3.73 mmol L�1. Individuals with an average wet weight of 20.4 ± 7.3 g were

kept in original aquaculture water until used for experiments. The animals were killed in

ice-water immediately prior to incubation experiments or gut dissection. Whole animals

were incubated in 100-mL glass bottles with 30 mL of 0.2-μm filtered aquaculture

water that was aerated before use via an airstone. The bottles were sealed gas-tight with

rubber stoppers. For incubating intact guts, freshly killed animals were dissected along

their dorsal side by scissors and the gut was carefully removed from the animals by

ethanol-rinsed forceps. Dissected complete guts (gut content + wall) were incubated in

6-mL exetainer vials (Labco, High Wycombe, UK) that contained 1 mL of aerated,

0.2-μm filtered aquaculture water. The headspace of all incubation vials was taken with

atmospheric air. In addition to these oxic incubations, dissected complete guts and gut

walls were also incubated under anoxic conditions in 6-mL exetainer vials that

139


contained 1 mL of N2-flushed, 0.2-μm filtered aquaculture water. As control, N2-flushed,

0.2-μm filtered aquaculture water was incubated in gas-tight 100-mL glass bottles. The

headspace of all anoxic incubations was flushed by dinitrogen gas for 5 to 10 min. For

each incubation assay, 4-15 replicates were run with 1 individual or dissected gut per

incubation vial. Incubations were conducted at an average temperature of 28 ± 2°C for

maximally 3 h. The incubation vials were placed on a shaker to enforce the equilibration

of N2O between water and headspace. Accumulation of N2O was followed by regularly

taking 1-mL headspace samples through the rubber stopper (every 15 to 60 min).

Analysis of N2O production by gas chromatography (GC 7890 Agilent Technologies)

and calculation of N2O emission rates were made as described in Heisterkamp et al.

(2010).

Rate of total denitrification in dissected guts

The potential total denitrification (i.e. the production of N2O+N2) of dissected guts was

measured with the acetylene inhibition technique (Sørensen 1978). Freshly dissected

guts were incubated in 6-mL exetainers with 1 mL of N2-flushed, 0.2-μm filtered

aquaculture water and were incubated in an atmosphere of 10% acetylene and 90%

dinitrogen gas. Sampling, analysis of N2O, and calculation of N2O production rates

were made as described above.

Microscale distribution of oxygen and pH in dissected guts

Microsensors for oxygen and pH measurements in dissected guts were constructed as

previously described (Schulthess et al. 1981, Revsbech 1989). The tip diameters of the

sensors were 10–20 μm. The sensors were calibrated before, during, and after the

measurements. Oxygen microsensors were calibrated in Ringer’s solution (Merck,

Germany) at 0 and 100% air-saturation by purging with dinitrogen gas and synthetic air,

respectively. The pH sensors were calibrated in standard solutions of pH 7.0 and 9.0.

Freshly dissected guts were fixed on an agarose bottom in a flow-cell, which was

continuously flushed with oxygenated Ringer’s solution (Stief & Eller 2006). Aided by

a dissection microscope, the microsensor tip was positioned at the outer surface of the

gut wall, which was then defined as depth zero. Vertical steady-state concentration

profiles were recorded through the gut, starting 1 mm above the gut surface in the

140


oxygenated Ringer’s solution and measuring in increments of 0.1 mm to maximal 3 mm

below the gut surface into the agarose bottom. Profiles were run in the fore-, mid-, and

hind gut at different degrees of gut filling. All measurements and calibrations were

made at 28°C.

Nitrous oxide concentration and production rate in aquaculture water

The in situ concentration of N2O in the aquaculture water was determined by filling

100 mL unfiltered aquaculture water into 125-mL gas-tight bottles (n = 10) that

contained 4 mL saturated HgCl2 to inhibit any biological activity. The bottles were

shaken for several hours for equilibration of N2O between water and headspace and

afterwards the N2O concentration in the headspace was analyzed by GC measurements.

The N2O concentration in the water was calculated after Weiss & Price (1980).

Furthermore, unfiltered aquaculture water was aerated via an airstone and 100 mL of the

oxygenated water was incubated in 100-mL glass bottles (n = 3) that were sealed with

rubber stoppers. The water was continuously stirred with a glass-coated magnetic

stirring bar. Water samples (3 mL) were taken every 20 min through the rubber stopper

and transferred into N2-flushed 6-mL exetainers. After 1 h of oxic incubation, the water

in the bottles was flushed with dinitrogen gas for 10 min, the bottles were sealed again

and the remaining headspace was purged with dinitrogen gas. After this oxic-anoxic

shift, water samples (3 mL) were taken in regular time intervals for a total incubation

period of 16 h. The headspace in the exetainers was analyzed for N2O by GC

measurements after equilibration of N2O between water and gas phase. Calculation of

N2O production rates were made as described in Heisterkamp et al. (2010).

Results and Discussion

In situ rates and origin of nitrous oxide emission from shrimp

Under in situ conditions (28°C, 10 mmol L�1 nitrate), the aquacultured shrimp

L. vannamei emitted N2O with a mean rate of 4.33 nmol individual�1 h�1, equivalent to

0.20 nmol g�1 wet weight h�1 (Figure 1). On an individual basis, this is the highest rate

141


recorded for any aquatic invertebrate so far and is much higher than the average N2O

emission rates of coastal and freshwater invertebrate species from natural habitats that

are 0.32 and 0.07 nmol ind.�1 h�1, respectively, at 21°C and in situ nitrate concentrations

(0-0.45 mmol L�1 nitrate, Stief et al. 2009, Heisterkamp et al. 2010). The in situ N2O

emission rate of L. vannamei determined in this study is 21% higher than the mean rate

measured for L. vannamei specimens of the same size, but obtained from a different

RAS and incubated at different conditions (21°C, 2 mmol L�1 nitrate) (Heisterkamp et

al. 2010). Microbial N2O production associated with L. vannamei may thus be

stimulated by temperature and/or nitrate as previously shown for terrestrial and

freshwater invertebrate species (Karsten & Drake 1997, Matthies et al. 1999, Stief et al.

2010, Stief & Schramm 2010). The temperature of 28°C is in the typical range of

rearing temperatures used for L. vannamei and the nitrate concentration was in the upper

range of nitrate concentration reported for intensive aquaculture systems (Burford et al.

2003, Holl et al. 2010). The in situ rate reported here might therefore represent a typical

in situ N2O emission rate of L. vannamei in aquaculture systems.

N2O

(n

mo

l in

div

idu

al-1

h-1

)

0

1

2

3

4

5

6

7

8

9

10

11

12

13oxic

anoxic

Complete gutWhole animal

Gut wallComplete gut

(+ acetylene)

Filtered aquaculture

water

Figure 1: Rates of N2O emission from the shrimp Litopenaeus vannamei and its dissected guts incubated in original 0.2-�m filtered aquaculture water at 28°C. Whole animals of L. vannamei were incubated under oxic conditions; dissected complete guts (content + wall) were incubated under oxic and anoxic conditions; and gut walls only were incubated under anoxic conditions. Dissected complete guts of L. vannamei were also incubated under anoxic conditions with 10% acetylene, which inhibits the last step of denitrification. The resulting N2O production indicates total denitrification. Filtered aquaculture incubated under anoxic conditions served as control (here, the N2O production rate is presented as nmol L�1 h�1). Means ± SD of n = 4 to 15 are shown.

142


The N2O emitted from the shrimp was mainly produced in the animal’s gut (Figure 1).

Dissected complete guts (i.e. gut content + gut wall) incubated under anoxic conditions

produced N2O at a rate equivalent to 84% of the total N2O emission rate of the whole

animal. In contrast, the gut walls only produced minute amounts of N2O under anoxic

conditions. It can be ruled out that the filtered aquaculture water that was added to the

incubation vials biased the results, as the negative control produced only trace amounts

of N2O (9 × 10�5 nmol mL�1 h�1). Hence, N2O production mainly took place in the gut

contents, which strongly indicates that N2O production in the shrimp gut is primarily

mediated by ingested microbes. Incubation of complete guts under oxic conditions

resulted in a N2O emission rate 16 times lower than that under anoxic conditions. These

findings suggest that N2O production associated with L. vannamei is mainly due to

denitrification by ingested bacteria in the anoxic gut of the animal. This is in accordance

with observations made for freshwater and terrestrial invertebrates where ingested

denitrifiers are the key players in N2O production (Ihssen et al. 2003, Horn et al. 2006,

Stief et al. 2009). The bacterial abundance in aquaculture water is generally high due to

the copious supply of inorganic and organic nutrients and can reach up to 3.9 × 108 cells

mL�1 in RAS (Burford et al. 2003, Browdy & Jory 2009, Holl et al. 2010). L. vannamei

mainly takes up particle-attached microorganisms by feeding on particulate organic

matter and uses these microorganisms as additional food source (Avnimelech 1999, De

Schryver et al. 2008). The N2O production measured in the gut suggests, however, that

at least part of the ingested microorganisms survive and remain or even become

metabolically active in the gut, including facultative denitrifying bacteria.

We therefore specifically tested for the denitrification potential of the shrimp gut. Under

anoxic conditions, the total denitrification rate of complete guts was on average

9.9 nmol ind.�1 h�1 (192 nmol g�1 wet weight h�1) compared to the mean net rate of N2O

production of 3.6 ind.�1 h�1 (70 nmol g�1 wet weight h�1) (Figure 1). The N2O/N2 ratio

of gut denitrification was thus on average 0.57, resulting in the mean N2O yield of

36.5% from total denitrification (N2O/N2O+N2). This value is in the same range as

observed for denitrification in the gut of freshwater insect larvae and terrestrial

earthworms and is much higher than the N2O yields of usually below 1% in aquatic

sediments and water columns (Seitzinger 1988, Drake & Horn 2007, Bange 2008, Stief

et al. 2009). Aside from true denitrifiers, also nitrate-reducing (NO3� � NO2

�) and

nitrate-ammonifying bacteria (NO3� � NH4

+) might have contributed to the N2O

143


production in the anoxic gut. However, these bacteria produce N2O as by-product at

much lower production rates than true denitrifiers (Drake & Horn 2007). Additionally,

earthworm guts were shown to exhibit only a minor capacity of nitrate ammonification

(Ihssen et al. 2003). Furthermore, it is very unlikely that nitrification contributed to the

N2O production under anoxic conditions, since anoxia generally precludes nitrification

activity. Under oxic conditions, the low N2O production rate might be due to

nitrification activity. However, as the gut is probably completely anoxic under in vivo

conditions (see below), the N2O emission rates of dissected guts measured under anoxic

conditions are very likely to reflect in vivo N2O emission rates and nitrification does not

contribute to N2O production under in vivo conditions. The stimulation of the N2O

emission rate of L. vannamei by temperature and nitrate concentration can be expected

to be controlled in the same way as reported for freshwater invertebrates (Stief et al.

2009). The temperature of 28°C is within the optimal range for L. vannamei (Wyban et

al. 1995, Ponce-Palafox et al. 1997) and for denitrifying bacteria (Klotz et al. 2011), and

is likely to promote high feeding rates of L. vannamei and high metabolic rates of

denitrifying bacteria. Both parameters have the potential to increase N2O emission from

L. vannamei via higher cell numbers of ingested bacteria and/or higher denitrification

rates inside the gut (Stief et al. 2009, Stief & Schramm 2010). Since shrimp feed on

water-soaked food particles and also drink water at a rate of ca. 10 μL g�1 wet weight

h�1 (Dall & Smith 1977), the nitrate concentration in the gut was probably also in the

millimolar range. Such high nitrate concentrations may not only stimulate the total

denitrification rate, but also specifically inhibit the N2O reductase and thus increase the

N2O yield of denitrification (Blackmer & Bremner 1978).

Factors controlling the nitrous oxide yield in gut denitrification

The high fraction of incomplete denitrification in the shrimp gut may result from the

inhibition of the N2O reductase by the presence of oxygen or by low pH values in the

animal’s gut (Zumft 1997, Bergaust et al. 2010). However, microsensor measurements

revealed that filled dissected guts of L. vannamei rapidly consumed oxygen that

diffused from the air-saturated Ringer’s solution into the gut, resulting in anoxic

conditions throughout almost the entire gut diameter (Figure 2 A). Even in empty guts,

the diffusion of oxygen from the air-saturated Ringer’s solution could not fully

oxygenate the entire gut, since the core of the gut remained hypoxic (Figure 2 B). No

144


Filled gut

Dis

tan

ce

(m

m)

-1,0

-0,5

0,0

0,5

1,0

1,5

2,0

2,5

3,0

fore gutmiddle guthind gut

Empty gut

Oxygen (µmol L-1)

0 25 50 75 100 125 150 175 200 225

Dis

tan

ce

(m

m)

-1,0

-0,5

0,0

0,5

1,0

1,5

2,0

2,5

3,0

pH

6,0 6,5 7,0 7,5 8,0 8,5 9,0

Dis

tan

ce

(m

m)

-1,0

-0,5

0,0

0,5

1,0

1,5

2,0

2,5

3,0

3,5

Filled gut

A

B

C

obvious variation in oxygen concentrations along the gut axis was observed in filled and

in empty guts (Figure 2 A+B).

Figure 2: Transversal micropro-files of oxygen and pH through dissected guts of Litopenaeus vannamei. Dissected guts were fixed on an agarose bottom in a flow-cell that was continuously supplied with air-saturated Ringer’s solution. Grey area indicates gut interior and dashed lines indicate gut walls. Upper and lower white areas indicate Ringer’s solution and agarose bottom, respectively. A: Oxygen concentration profiles through filled guts at fore, middle, and hind position. B: Oxygen concentration profiles through empty guts at fore, middle, and hind position. For A and B, single representative profiles are shown. C: pH profile (mean ± SD, n = 6) through filled guts calculated from pH profiles measured in fore-, mid-, and hind guts.

Under in vivo conditions, the oxygen flux into the gut is probably much lower than after

dissection. The hemolymph surrounding the shrimp’s gut has typically a much lower

oxygen concentration (ca. 2-3 �mol L�1 O2) than the air-saturated Ringer’s solution

145


(Chen & Cheng 1995, Lin & Chen 2001). Therefore, it can be assumed that under in

vivo conditions the entire gut is anoxic even when it is not completely filled. Hypoxic

conditions may only prevail in empty guts. The number of bacteria in empty guts is

much lower than in filled guts, which is also supported by very low N2O emission rates

(Figure 1). The high N2O yield of denitrification in the gut of L. vannamei is therefore

probably not explained by the inhibition of the N2O reductase by oxygen, since

gradients of oxygen along the gut radius or the gut axis are unlikely to occur in filled

denitrifying guts. Furthermore, it is unlikely that the N2O reductase is inhibited by pH,

given the fact that the pH in the gut is 7.6-7.8 and thus favourable for complete

denitrification and no greater changes in pH were observed along the gut axis

(Figure 2 C). A pH of below 6.5-6.8 was reported to be critical for the functioning of

the N2O reductase and a pH of 6.0 almost completely inhibits the reduction of N2O to

N2 (Baumann et al. 1997, Bergaust et al. 2010).

The high N2O yield of denitrification in the gut of L. vannamei may result from the

delayed expression of the N2O reductase. This scenario seems likely because of the very

short gut residence time of ingested bacteria in L. vannamei of only about 1 h (Beseres

et al. 2005). When water samples (and the microorganisms contained therein) from the

rearing tank were experimentally transferred from oxic to anoxic conditions, N2O

production was immediately stimulated and increased by a factor of 4.5 to a rate of

0.92 ± 0.18 nmol L�1 h�1 compared to the very low N2O production rate of 0.20 ± 0.28

nmol L�1 h�1 under oxic conditions (Figure 3).

This indicates that the oxic-anoxic shift makes facultative anaerobic bacteria switch

metabolically from aerobic respiration to denitrification with at first unbalanced enzyme

activities. Immediately after the oxic-anoxic shift, an accumulation of N2O was

observed, which started to disappear after a period of at least 2.5 to 3.5 h, indicating the

incipient activity of the N2O reductase (Figure 3). This strongly suggests that facultative

denitrifying bacteria that are ingested by L. vannamei get activated in the anoxic gut of

the shrimp, but the gut passage time of approximately 1 h does not allow them to

balance the expression of the four denitrification genes. It seems that the induction of

the N2O reductase is initially weaker or lags behind that of the other denitrifying

enzymes, which consequently causes high N2O yields of denitrification during the short

gut passage time. Similarly high N2O yields of gut denitrification are reported for the

146


larvae of the insect Chironomus plumosus that has a gut passage time of 2 to 3 h (Stief

et al. 2009). The gut passage time seems thus to be an important factor regulating the

N2O yield of denitrification in the gut of invertebrates. The transient accumulation of

N2O after shifts from oxic to anoxic conditions was also shown for several pure cultures

of denitrifiers (Baumann et al. 1996, Otte et al. 1996, Kester et al. 1997, Bergaust et al.

2008). The time required to balance denitrification enzyme activity and perform

complete denitrification after the oxic-anoxic shift varies with species and culture

conditions. The community composition of denitrifiers ingested by L. vannamei might

therefore play an important role. For some species of denitrifying bacteria, gut passage

times of 1-3 h might be too short to express the full set of genes, while other species

might switch to complete denitrification within a few minutes. Furthermore, the

denitrifying community may comprise many species that are not even capable of

complete denitrification because they lack the gene encoding the N2O reductase. It is

estimated that N2O-respiring taxa make up only 10-15% of all known denitrifying taxa

(Zumft & Kroneck 2007). N2O-reducing taxa might be underrepresented in the highly

nitrate-enriched RAS, since the reduction of N2O is inhibited by high nitrate

concentrations (Blackmer & Bremner 1978, Richardson et al. 2009).

Time (h)

0 2 4 6 8 10 12 14 160,0 0,2 0,4 0,6 0,8 1,0

N2O

(n

mo

l)

0,0

0,1

0,2

0,3

0,4

0,5

0,6

0,7

0,8

0,9

1,0oxic anoxic

Figure 3: Induction of N2O production in aquaculture water by anoxia. Left white panel shows N2O emission in three replicate incubations of aquaculture water under oxic conditions for a period of 1 h. Shaded area indicates shift from oxic to anoxic conditions. Right grey panel shows N2O emission from aquaculture water in the same three replicates under anoxic conditions during an incubation period of 16 h.

147


Relevance of nitrous oxide emission from L. vannamei

re. Aquaculture farming of

. vannamei thus represents a source of atmospheric N2O.

r, and no dilution (the

ater exchange rate in the RAS was only about 1-3% per day).

The water in the rearing tanks of L. vannamei contained a mean N2O concentration of

160 ± 17 nmol L�1, which corresponds to a supersaturation of 2390 ± 257% compared

to the atmospheric equilibrium concentration of 6.67 nmol L�1. The permanent aeration

of the shallow rearing tanks leads to continuous water movement and air stripping,

which probably results in high N2O fluxes to the atmosphe

L

Nitrous oxide emission by L. vannamei accounted for 2.17 nmol L�1 h�1 at a density of

0.5 individuals L�1, which is equivalent to 100 individuals m�2 and represents a

common stocking density in RAS (Browdy & Jory 2009, Krummenauer et al. 2011).

The N2O production in the aquaculture water under oxic conditions was

0.2 nmol L�1 h�1 and thus 10 times lower than by the shrimp. Under anoxic conditions,

the initial rate of N2O production in the aquaculture water increased compared to the

rate under oxic conditions, but was still less than half of the shrimp-associated N2O

production rate. Hence, denitrification in the aquaculture water under non-steady state

conditions obviously produced more N2O than nitrification under completely oxic

conditions. In any case, the microorganisms in the small gut of the shrimp produced

more N2O than the microorganisms contained in 1 Liter of aquaculture water. These

findings suggest that the supersaturation of the water is to a considerable extent due to

microbial N2O production associated with L. vannamei. The steady-state concentration

of 160 nmol L�1 N2O in the water of the rearing tank could theoretically be built up

solely from N2O emission of L. vannamei within 3 days, assuming no release of N2O to

the atmosphere, no N2O consumption in the aquaculture wate

w

However, the N2O production rate of the aquaculture water measured under fully oxic

conditions is probably an underestimate of in situ N2O emission from the rearing tanks.

High net N2O production rates are typically observed under hypoxic conditions, since

the N2O yield of both nitrification and denitrification increases under suboptimal

conditions (Goreau et al. 1980, Bonin & Raymond 1990). Oxygen concentrations in

aquaculture systems can vary greatly and are often lower than at air saturation due to

high respiration rates in the densely colonized rearing tanks (Cuzon et al. 2004). Higher

148


N2O production rates than measured under fully oxic conditions might therefore prevail

under hypoxic conditions in the water column and especially in the low-oxygen

microsites of organic particles. Additionally, the rates of nitrification and denitrification

(and concomitant N2O production) vary greatly between the different compartments of

aquaculture systems. About 75-100% of the volume of the aquaculture water is pumped

through nitrification biofilters hourly. In these biofilters, nitrification rates and possibly

also N2O production rates are much higher than in the water from the rearing tanks due

to a large nitrifying biomass. N2O production rates of 0.5-4 μmol L�1 h�1 were reported

for a nitrifying biofilter in a wastewater treatment plant, although the N2O yield from

nitrification was only 0.4% (Tallec et al. 2006). Whether N2O production rates in the

biofilters of the RAS investigated here are similarly high, still needs to be investigated.

In addition, N2O production might also take place in organic food particles and in

floating feces that are probably characterized by steep oxygen gradients and an anoxic

core. Feces might be hotspots of N2O production, since they are enriched in active

denitrifiers that once more experience a sudden shift in oxygen concentration when

being voided into the oxic rearing water. Taken together, the supersaturation of N2O in

the aquaculture water is probably caused by many different N2O sources. The high in

situ N2O emission rate of L. vannamei and its high density in the RAS, however,

suggests that the shrimp significantly contributes to the N2O supersaturation of the

quaculture water and thus to emission of N2O to the atmosphere.

onclusion and Outlook

a

C

The gut of the aquacultured shrimp L. vannamei constitutes an anoxic microsite of

massive N2O production by incomplete denitrification of ingested bacteria. The high

N2O yield of gut denitrification is apparently due to the gut passage time of only 1 h,

which may be too short for many of the ingested denitrifiers to establish the complete

denitrification pathway. There are no indications that the N2O reductase of the ingested

denitrifiers is inhibited by unfavourable conditions in the gut of L. vannamei. Further

investigations are underway that assess the relative abundance and expression levels of

denitrification genes and the microbial community composition in different sections of

the gut and the water column. This is expected to shed light upon the fate and activity of

149


ingested facultative denitrifiers in the animal’s gut. The in situ N2O emission rates of

L. vannamei suggest that the shrimp significantly contributes to the overall N2O

emission from aquaculture farms. A direct comparison of N2O production in the

different compartments of aquaculture systems, including the aquaculture water,

suspended feces and food pellets, animal guts, and the integrated biofilters, would be

highly desirable. In the light of the fast-growing aquaculture industry, especially of

penaeid shrimp species like L. vannamei, it is important to gain insight into the

echanisms and controlling factors of N2O production in aquacultures.

m

150


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154

Chapter 6

The polychaete Hediste diversicolor is one of the most important bioturbating species in temperate coastal marine sediments that influences nitrogen cycling by changing physico-chemical conditions in its surrounding.

155

Indirect control of the intracellular nitrate poolof intertidal sediment by the

polychaete Hediste diversicolor

Ines M. Heisterkamp, Anja Kamp, Angela T. Schramm,

Dirk de Beer, and Peter Stief

Max Planck Institute for Marine Microbiology, Microsensor Group,


Marine Ecology Progress Series 445:181-192, 2012

157

Chapter 6 Intracellular nitrate in intertidal sediment

Abstract

In an intertidal flat of the German Wadden Sea, a large sedimentary pool of intracellular

nitrate was discovered that by far exceeded the pool of nitrate that was freely dissolved

in the porewater. Intracellular nitrate was even present deep in anoxic sediment layers

where it might be used for anaerobic respiration processes. The origin and some of the

ecological controls of this intracellular nitrate pool were investigated in a laboratory

experiment. Sediment microcosms were set up with and without the abundant

polychaete Hediste diversicolor that is known to stimulate nitrate production by

microbial nitrification in the sediment. Additional treatments were amended with

ammonium to mimic ammonium excretion by the worms or with allylthiourea (ATU) to

inhibit nitrification by sediment bacteria. H. diversicolor and ammonium increased,

while ATU decreased the intracellular nitrate pool in the sediment. Microsensor profiles

of porewater nitrate showed that bacterial nitrification was enhanced by worms and

ammonium addition. Thus, nitrification formed an important nitrate supply for the

intracellular nitrate pool in the sediment. The vertical distribution of intracellular nitrate

matched that of the photopigments chlorophyll a and fucoxanthin, strongly suggesting

that diatoms were the main nitrate-storing organisms. Intracellular nitrate formation is

thus stimulated by the interaction of phylogenetically distant groups of organisms:

Worms enhance nitrification by feeding on particulate organic matter, excreting

ammonium, and oxygenating the sediment. Bacteria oxidise ammonium to nitrate in

oxic sediment layers, and diatoms store nitrate intracellularly.

158


Introduction

Several phylogenetically distant groups of sediment microorganisms are able to store

nitrate in their cells. Large sulphur bacteria (Schulz & Jørgensen 2001), foraminifera

(Risgaard-Petersen et al. 2006, Pina-Ochoa et al. 2010), and microalgae (Garcia-

Robledo et al. 2010, Kamp et al. 2011) store nitrate at millimolar concentrations, while

in their direct environment porewater nitrate is available only at micromolar

concentrations. Thus, the uptake of nitrate into the cell occurs against a steep

concentration gradient and costs metabolic energy (Høgslund et al. 2008). Storage of

nitrate is of obvious advantage in environments with fluctuating nutrient concentrations.

Also, intracellular nitrate is known to be used for respiration in anoxic sediment layers.

The high nitrate storage capacity of large sulphur bacteria enables them to survive long

periods of anoxia when intracellular nitrate is respired to ammonium (Preisler et al.

2007, Høgslund et al. 2009). Respiratory use of intracellular nitrate has recently also

been shown for microeukaryotes such as foraminifera (Risgaard-Petersen et al. 2006)

and diatoms (Kamp et al. 2011). At the oxic sediment surface, however, diatoms and

other microalgae use dissolved inorganic nitrogen (DIN) and probably also intracellular

nitrate for nitrogen assimilation (Lomas & Glibert 2000, Sundbäck & Miles 2000).

The ability to store nitrate intracellularly may be particularly advantageous in intertidal

flats which are very dynamic ecosystems. Benthic organisms have to cope with frequent

changes in the availability of, e.g., light, oxygen, and nutrients due to tidal and diurnal

rhythms. Another source of perturbation in intertidal flats is the presence of macrofauna

that reworks large amounts of sediment and the microorganisms therein (e.g., Bouchet

et al. 2009). Some polychaetes construct deep-reaching burrows and enhance solute

exchange between sediment and the water column due to their ventilation activity

(Kristensen 2001). Many species of intertidal macrofauna feed on sediment

microorganisms and thereby decrease microbial populations or keep them in the

exponential growth phase (Herman et al. 2000, Blanchard et al. 2001). Under such

transient conditions, the nitrate storage capacity awards sediment microorganisms with

the steady availability of a key nutrient and an energetically favourable electron

acceptor. Nitrate-storing microorganisms may thereby gain a competitive advantage

over sediment bacteria that lack the ability to store nitrate intracellularly.

159


Nitrate is supplied to intertidal sediments via the water column or is produced by

microbial nitrification at the oxic sediment surface. Nitrate from nitrification diffuses

both into the water column and towards anoxic layers in the sediment where it can be

anaerobically respired to dinitrogen gas by microbial denitrification. In intertidal

sediments, the coupling of nitrification and denitrification can be relatively loose

(Jensen et al. 1996). This implies that either denitrification has a substantial nitrate

source other than nitrification (e.g., the water column) or that much of the nitrate

produced by nitrification does not end up as dinitrogen produced by denitrification.

Nitrate-storing microorganisms take up nitrate from the water column or from the

nitrification layer of the sediment surface (Sayama 2001). The sedimentary intracellular

nitrate pool might be controlled by the rates of nitrogen mineralisation and nitrification

in the sediment. Both processes are stimulated by the oxygenation of the sediment by

tidal currents and by ventilation of macrofaunal burrows (Kristensen 2001, Nielsen et

al. 2004, de Beer et al. 2005). Additionally, some macrofauna species enrich the

sediment with organic matter due to their feeding activities (Christensen et al. 2000),

but also digest organic matter in their gut, which further enhances mineralisation and

ammonium regeneration (Gardner et al. 1993).

In an intertidal flat of the German Wadden Sea, a snapshot measurement revealed a

large pool of intracellular nitrate that reached deep into the sediment, well below the

very thin photosynthetic layer (de Beer et al. 2005). The sediment was densely

populated by diatoms, but also by the burrowing polychaete Hediste diversicolor.

Hence, in a laboratory microcosm experiment, the hypothesis was tested that the

presence of H. diversicolor in intertidal sediment increases the nitrate supply and

thereby the size of the sedimentary intracellular nitrate pool via stimulation of

nitrification. As experimental treatments served (1) sediment without worms,

(2) sediment with worms, (3) sediment without worms, but amended with ammonium to

mimic the worms’ ammonium excretion, and (4) sediment with worms, but amended

with the nitrification inhibitor allylthiourea.

160



Origin of sediment and animals

Sediment was collected in the intertidal flat near Dorum-Neufeld in the German

Wadden Sea (53°45'N, 8°21'E). This site is characterised by mixed sediment

(sand/mud) with low porewater sulphide concentrations (Jahn & Theede 1997) and high

densities of epifauna (e.g., the snails Hydrobia ulvae and Littorina littorea) and infauna

(e.g., the polychaetes Arenicola marina and Hediste diversicolor). Sediment from the

top 25 cm was sieved through a 0.5 mm screen to remove macrofauna and shell debris.

It was then frozen at �20°C for 30 h to kill macrofauna juveniles that had passed

through the sieve. The defaunated and homogenised sediment was added to

4 recirculating flow-through microcosms (30 cm long × 20 cm wide × 10 cm high). A

thin layer of unfrozen and 180 �m sieved sediment was evenly distributed on the

sediment surface to inoculate the sediments with living microalgae. After the sediment

had settled, aerated seawater from the North Sea diluted to the in situ salinity of 22 was

continuously directed over the sediment surface. Each microcosm was continuously

supplied from its own 50 L seawater reservoir at a flow rate of 3 L min�1 throughout the

experiment. To allow the growth of microalgae on the sediment surface and the

formation of the typical redox stratification in the sediment, the flow-through

microcosms were illuminated from above by a neon daylight lamp (50 μmol photons

m�2 s�1 light intensity at the sediment surface) at a 16 h light to 8 h dark cycle and left

untouched for 10 days. The incubation temperature was 22°C, which was at the upper

end of temperatures reached at the collection site during summer when large infauna is

abundant and exhibits high foraging, burrowing, and ventilation activities. The

polychaete Hediste diversicolor (O.F. Müller) was freshly collected in the intertidal flat

near Dorum-Neufeld by digging up the sediment with a spade to a depth of

approximately 25 cm and searching it through by hand. On the day of collection,

30 individuals of 250 to 300 mg wet weight were added to 2 of the 4 microcosms,

which corresponded to a density of 420 ind. m�2.

161


Experimental design

The experiment comprised four treatments (one in each microcosm): A) Sediment

without H. diversicolor (Control), B) Sediment colonised by H. diversicolor (Hediste),

C) Sediment not colonised by H. diversicolor, and overlain with ammonium-enriched

water (Ammonium), and D) Sediment colonised by H. diversicolor, and overlain by

allylthiourea-treated water (Hediste + ATU). Allylthiourea, an inhibitor of microbial

ammonia oxidation (Hall 1984), was added to the seawater at a final concentration of

100 μmol L�1 on the day the animals were added. Three days later, NH4Cl was added to

the seawater of treatment C) to a final concentration of 50 μmol L�1 NH4+; 16 days later

it was replenished because the concentration had dropped to less than 2 μmol L�1 NH4+.

Water samples from the four microcosms were taken every two days during the course

of the experiment and stored at -20°C until ammonium was analysed by flow-injection

(Hall & Aller 1992) and nitrate was analysed using the VCl3 reduction method (Braman

& Hendrix 1989) with a chemiluminescence detector (CLD 86 S NO/NOx-Analyser,

Eco Physics, Switzerland). Microsensor measurements were started 10 days after the

animals were added and were completed within 11 days. Afterwards, sediment cores

were taken for the analysis of intracellular nitrate and photopigments.

Intracellular nitrate

In the intertidal flat near Dorum-Neufeld (53°45'N, 8°21'E) that was densely colonised

by H. diversicolor, four randomly selected sediment cores with an inner diameter of

3.6 cm were taken at low tide. One sediment core was used for measuring porewater

nitrate concentration by microsensor measurements (see ‘Microsensor measurements’

below). The other three sediment cores were sliced at 0.2-cm intervals for the upper

1 cm and at 1-cm intervals to a total depth of 15 cm. Care was taken to remove

macrofauna from each slice with forceps. The sediment slices were frozen at -20°C until

used for nitrate extraction with the freeze-and-thaw technique (Lomstein et al. 1990).

For the extraction, 1 mL Milli Q water was added to the 0.2-cm sediment slices (upper

10 mm) and 3 mL Milli Q water to the 1-cm sediment slices (1-15 cm). Samples were

vigorously shaken, frozen in liquid nitrogen, and heated in the water bath (90°C) three

times for 10 min each to physically break up large microbial cells and thereby release

intracellular nitrate. The concentration of total nitrate (porewater nitrate plus extracted

162


nitrate) in the supernatant of the sediment slurries was measured using the VCl3

reduction method (Braman & Hendrix 1989). The intracellular nitrate concentration

(expressed in nmol cm�3 sediment) was calculated by subtracting the porewater nitrate

concentration from the total nitrate concentration.

In the microcosm experiment, four randomly selected sediment cores with an inner

diameter of 2.5 cm were taken from each microcosm and sliced at 0.2-cm intervals for

the upper 1 cm and at 1-cm intervals to a total depth of 5 cm. One half of each sediment

slice was used for intracellular nitrate analysis and was frozen at -20°C, the other half

was used for pigment analysis and was frozen at -80°C. Extraction and analysis of

intracellular nitrate were made as described above.

Photopigments

Sliced sediment from the laboratory microcosms was defrosted and each slice was

incubated with 10 mL 90% acetone (Sigma-Aldrich, Switzerland) on a rotary shaker at

4°C over night. After centrifugation for 10 min at 3700 g at 0°C, the supernatants were

filtered (Acrodisc® CR 4 mm Syringe Filter with 0.45 μm Versapor® Membrane,

Gelman Laboratory) and filled into HPLC-vials. Samples were always kept in the dark.

Extracted pigments were separated by means of HPLC (Waters 2695, U.S.A.) and

analysed by a photodiode array detector (Waters 996, U.S.A.). The HPLC column

(Reprosil, 350 × 4.6 mm, Dr. Maisch, Germany) was heated to 25°C, while the samples

were kept at 4°C during measurements. Pigments of each sample were separated by

three different eluents (methanol:ammonium acetate (80:20), acetonitrile 90% and ethyl

acetate (100%), flow rate 1 mL min�1) the mixing ratio of which changed gradually

during each 24 min run. Peaks were integrated with the software Millenium32 and

chlorophyll a and fucoxanthin peaks were identified according to their specific retention

time and absorption spectrum. Calibrations were performed by using 1:5, 1:10, 1:20 and

1:40 dilutions of a chlorophyll a stock solution (1.963 mg L�1, DHI, Denmark) and a

fucoxanthin stock solution (1.075 mg L�1, DHI, Denmark).

163


Microsensor measurements

Oxygen and NOx microsensors were constructed as described by Revsbech (1989) and

Larsen et al. (1997), respectively. The sensors were calibrated before and after each

series of 4-6 profiles in sterile seawater equilibrated to 22°C. Oxygen microsensors

were calibrated at 0 and 100% air saturation by flushing the seawater with either

dinitrogen gas or synthetic air. NOx microsensors were calibrated by adding aliquots of

a 10 mmol L�1 stock solution of NaNO3 to a known volume of seawater to arrive at

nominal nitrate concentrations of 0, 20, 40, 60, 80, and 100 μmol L�1. The calibration

curve was corrected for the natural concentration of nitrate in the seawater that was

determined with the VCl3 reduction method. The oxygen and NOx microsensors were

simultaneously used in a measuring set-up as described by Stief & de Beer (2002). At

least 4 and up to 14 vertical steady-state concentration profiles were recorded in each

flow-through microcosm from 0.3 cm above to 0.6 cm below the sediment surface in

increments of 0.025 cm. Positions of the profiles were randomly selected, but a

minimum distance of 2 cm to burrow openings at the sediment surface was held in the

two Hediste treatments. In the natural sediment core from the intertidal flat, profiling

with the NOx microsensor was done down to a depth of 2 cm in the laboratory within

2 h of collection. Profiles were recorded at three randomly selected spots of the

sediment surface.

The NOx profiles were interpreted as porewater nitrate profiles, assuming that nitrite

and nitrous oxide (N2O) concentrations in the sediments were negligible. For

calculating the depth-integrated nitrate content (see ‘Depth integration of data and

statistical analysis’), the concentration values in μmol L�1 porewater were converted to

concentration values in nmol cm�3 sediment by multiplication with the average

sediment porosity of 0.41. Local volumetric net nitrate production rates were calculated

from the curvature of the steady state NOx concentration profiles by diffusion-reaction

modeling (Bungay et al. 1969, Berner 1980). The effective diffusion coefficient of

nitrate at depth x in the sediment was calculated as Ds(x) = D0 × ��ln (�2))

(Boudreau 1996) with D0 as the diffusion coefficient of nitrate in seawater and � as the

sediment porosity. D0 of nitrate in seawater was taken as 1.75 × 10�5 cm2 s�1 at 22°C

(Li & Gregory 1974). � was determined as the volumetric water content of the

164


sediment, which corresponded to the weight loss of sediment slices of known wet

volume after drying at 60°C for 3 days.

Depth-integration of data and statistical analysis

Depth-integrated values of photopigments, intracellular nitrate (expressed per μg

chlorophyll a), porewater nitrate, and local nitrate production rates were calculated from

the respective vertical profiles. Depth-integrated contents of chlorophyll a, intracellular

nitrate, and porewater nitrate were calculated by adding up the average concentration

values of every depth interval of the vertical profile multiplied by the individual

thickness of each depth interval. Depth-integrated net nitrate production rates were

calculated by adding up the local production rates multiplied by the thickness of each

depth interval of the production-consumption profiles.

The depth-integrated contents of chlorophyll a, intracellular nitrate (expressed per μg

chlorophyll a), porewater nitrate as well as the depth-integrated net nitrate production

rates were compared between the four treatments. One-way ANOVAs were run for each

variable after confirming normality and homogeneity of variance of the data. If the null

hypothesis was rejected, the Waller-Duncan Post-hoc test was used for pairwise

comparisons. This test is based on Bayesian principles and uses the harmonic mean of

different samples sizes. For the depth-integrated contents of intracellular nitrate, an

additional T-test was run for the pairwise comparison between the treatments Control

and Hediste. All statistical analyses were carried out with the program SPSS Version

11.

Results

Sedimentary pool of intracellular nitrate

Under in situ conditions, the sedimentary pool of intracellular nitrate showed a peak

reaching from 0 to 5 cm depth with a maximum concentration of 11.7 nmol cm�3

sediment (Fig. 1). Below 5 cm, the intracellular nitrate concentration was relatively

165


constant around 2.5 nmol cm�3 sediment to the depth of 15 cm (Fig. 1). In contrast, the

porewater nitrate concentration was highest at the sediment-water interface (up to

4.1 nmol cm�3 sediment) and decreased rapidly to 0 within the upper 1.5 cm of the

sediment (Fig. 1).

Figure 1: In situ distribution of intracellular and porewater nitrate in intertidal sediment densely colonised by diatoms and burrowing macro-fauna. Both intracellular nitrate and porewater nitrate concentrations are given in nmol cm�3 sediment. Means ± SD of n = 3 replicate profiles are shown.

NO

3

- (nmol cm

-3 sediment)

0 5 10 15 20

Dep

th (

cm

)

0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

15

Porewater

Intracellular

In the microcosms, the sedimentary pools of intracellular nitrate showed the highest

concentrations at the surface and then decreased gradually with depth in all treatments

(Fig. 2A-D). Intracellular nitrate concentrations were generally higher and also extended

to greater depth in the Hediste and Ammonium treatments than in the Control and

Hediste + ATU treatments. The highest concentration of 71 nmol cm�3 sediment was

observed in the Ammonium treatment (Fig. 2C).

166


0

1

2

3

4

5

De

pth

(cm

)

0

1

2

3

4

50

1

2

3

4

5

Intracellular NO3

-

(nmol cm-3

sediment)

0 15 30 45 60 75 90

0

1

2

3

4

5

Control

Hediste

Ammonium

Photopigments

(�g cm-3

sediment)

0 3 6 9 12 15 18

Hediste + ATU

Chlorophyll a

Fucoxanthin

A

B

C

D

E

F

G

H

Figure 2: Vertical distribution of (A-D) intracellular nitrate and (E-H) photo-pigments in coastal marine sediment incubated in laboratory flow-through microcosms for 3 weeks. (A, E) Control sediment without H. diversicolor, (B, F) Sediment colonised by H. diversicolor, (C, G) sediment without H. diversi-color, overlain with ammonium-enriched water, (D, H) sediment colonised by H. diversicolor, overlain by allylthiourea (ATU)-treated water. Means ± SD of n = 4 replicate cores.

While in many of the sediment layers intracellular nitrate was detected, it was totally

absent from other layers. In contrast, organic matter was homogenously distributed in

all sediment layers because the sediment was thoroughly homogenised before it was

filled into the microcosms. From this we concluded that organic nitrogen compounds

that were potentially extracted and degraded by the extreme temperature changes were

not converted to nitrate, which would have produced false-positive results.

167


Chlorophyll a (µg cm-3

)

0 2 4 6 8 10 12

Intr

acellu

lar

NO

3

- (n

mo

l cm

-3)

0

10

20

30

40

50

R2 = 0.934

A)

Fucoxanthin (µg cm-3

)

0 1 2 3 4 5 6

Intr

acellu

lar

NO

3

- (n

mo

l cm

-3)

0

10

20

30

40

50

R2 = 0.955

B)

Photopigment distribution in the sediment

The vertical distributions of marker pigments of diatoms, chlorophyll a and

fucoxanthin, were similar in all treatments with the highest pigment concentrations at

the sediment surface and a gradual decrease down to 5 cm depth (Fig. 2E-H).

Fucoxanthin was present in all sediment slices, indicating the presence of viable

diatoms even in relatively deep sediment layers. In the upper 2 mm of the sediment, the

fucoxanthin-to-chlorophyll a ratio was particularly high (i.e., 0.4-0.6) and suggested

that the photosynthetically active community was dominated by diatoms (Lucas &

Holligan 1999), which was also confirmed by qualitative microscopic examination.

Average concentrations of the two photopigments within each of the 9 sediment layers

were linearly correlated with average concentrations of intracellular nitrate (Fig. 3).

Figure 3: Correlation of intracellular nitrate with A) chlorophyll a and B) fucoxanthin in sediment cores taken from the 4 laboratory microcosms. Nitrate and pigment contents were averaged for each of the 9 sediment layers analysed (compare Fig. 2). Error bars give SE for each sediment layer. R2 is Pearson’s coefficient for linear correlations.

168


Porewater concentrations of oxygen and nitrate

The oxygen profiles were identical in all treatments, except in the sediment of the

Hediste + ATU treatment, where oxygen penetrated deeper. Diffusion-reaction

modeling revealed no significant net oxygen production due to microalgal

photosynthesis at the sediment surface at which the light intensity was 50 μmol photons

m�2 s�1 (data not shown). The nitrate concentration profiles showed surface peaks

indicative of nitrate production by nitrification in all treatments, except the

Hediste + ATU treatment (Fig. 4A-D). Diffusion-reaction modeling revealed net

production of nitrate in the oxic sediment layer of the Control, Hediste, and Ammonium

treatments (Fig. 4E-G), but not in the Hediste + ATU treatment (Fig. 4H). Below the

oxic surface layer, net nitrate consumption by bacteria and/or microalgae occurred,

which did not show any significant differences between all non-inhibited treatments, but

which was very low in the Hediste + ATU treatment (Fig. 4E-H). It should be noted that

net nitrate production and consumption might also have occurred inside the Hediste

burrows, but this microbial activity was not measured with the microsensor approach.

The nitrate concentration in the water column of the Hediste + ATU treatment was

lower than in the natural North Sea water due to the inhibition of nitrification by ATU

(Fig. 4A-D, Tab. 1). The ammonium concentration in the water column was highest in

the ATU-inhibited treatment, intermediate in the Ammonium treatment and lowest in

the Control and Hediste treatments (Tab. 1).

Table 1: Mean ammonium and nitrate concentrations in the water column of the four sediment microcosms over the incubation period of three weeks. Treatments are described in the legend of Fig. 2. ATU: allylthiourea

Treatment NH4+ [μmol L�1] (± SD) NO3

� [μmol L�1] (± SD)

Control 1.2 (0.7) 34.1 (9.8)

Hediste 2.7 (2.2) 44.8 (7.9)

Ammonium 11.2 (8.9) 46.2 (17.6)

Hediste + ATU 33.5 (6.1) 14.0 (5.4)

169


-0,4

-0,2

0,0

0,2

0,4

0,6

NO3

- (nmol mL

-1)

0 25 50 75 100 125

NO3

- rate (nmol cm

-3 h

-1)

0 250 500 750

Nitrate

Oxygen

-0,4

-0,2

0,0

0,2

0,4

0,6

De

pth

(cm

)

-0,4

-0,2

0,0

0,2

0,4

0,6

O2 (nmol mL

-1)

0 50 100 150 200 250

-0,4

-0,2

0,0

0,2

0,4

0,6

Control

Hediste

monium

+ ATU

E

F

G

H

Am

Hediste

A

B

C

D

Figure 4: (A-D) Vertical microprofiles of porewater oxygen and nitrate in laboratory sediment microcosms and (E-H) nitrate conversion rates derived from the microprofiles. Treatments as described for Fig. 2. Dashed line indicates the sediment-water interface. Positive and negative rates correspond to net production and consumption of nitrate, respectively. Means ± SD of n = 4 to 14 replicate profiles are shown.

170


Co

ntr

ol

He

dis

te

Am

mo

niu

m

He

dis

te +

AT

U

Chla

(µg c

m-2

)

0

4

8

12

16

20

NO

3

- per � g

Chl a

(nm

ol cm

-2)

0

2

4

6

8

10

NO

3

- (n

mo

l cm

-2)

0

5

10

15

20

D) Chlorophyll a

B) Porewater NO3

-

A) Intracellular NO3

-

aa

a

a

a

b

b

c

a

a

b

a

NO

3

- (n

mo

l cm

-2 h

-1)

0

10

20

30

40

50

60 C) NO3

- Production

a

b

a

c

Depth-integrated contents of nitrate and photopigments

Significant differences between the four treatments were found with respect to the

depth-integrated contents of intracellular nitrate (expressed per μg chlorophyll a),

porewater nitrate, and net nitrate production rates (ANOVA: F3,12 = 19.7, p < 0.001,

F3,25 = 12.8, p < 0.001, and F3,25 = 12.1, p < 0.001, respectively) (Fig. 5A-C). In

contrast, no significant differences between the

four treatments were found with respect to the

depth-integrated contents of chlorophyll a

(ANOVA: F3,12 = 0.8, p = 0.512, Fig. 5D) and

fucoxanthin (ANOVA: F3,12 = 2.6, p = 0.097, data

not shown). The porewater nitrate contents and the

net nitrate production rate were significantly higher

in the Hediste than in the Control treatment

(Waller-Duncan Post-hoc test, Fig. 5B+C). Also

the intracellular nitrate content (expressed per μg

chlorophyll a) was significantly higher in the

Hediste than in the Control treatment, but only in a

pairwise comparison that excluded the Ammonium

treatment with its extraordinarily high average

intracellular nitrate content (Student’s T-test:

T6 = -2.8, p < 0.05, Fig. 5A).

Figure 5: Depth-integrated A) intracellular nitrate (expressed per μg chlorophyll a), B) porewater nitrate, C) net nitrate production, and D) chlorophyll a. Treatments are described in Fig.2. Means + SD of n = 3-14 replicate measurements are shown. Treatments with different lower case letters have significantly different means (ANOVA, Waller-Duncan Post-hoc test).

171


In the Ammonium treatment mimicking ammonium excretion by H. diversicolor, the

intracellular and porewater nitrate contents (but not nitrate production and chlorophyll

a) were significantly higher than in the Control treatment (Waller-Duncan Post-hoc test,

Fig. 5A-D). In the Hediste + ATU treatment, the intracellular and porewater nitrate

contents as well as the net nitrate production rate (but not chlorophyll a) were

significantly lower than in the Hediste and Ammonium treatments in which ammonium

was abundant and could be nitrified (Waller-Duncan Post-hoc test, Fig. 5A-D).

The variability of depth-integrated profiles within each treatment was high for the

punctual oxygen and nitrate microsensor profiles (i.e., coefficients of variation

(CV) = 29-39 and 31-40%, respectively) and low for chlorophyll a and fucoxanthin

profiles that were analysed in sediment cores that integrated over ca. 5 cm2 of sediment

surface (i.e., CV = 2-22 and 7-15%, respectively). Intracellular nitrate that was also

analysed in sediment cores took an intermediate position with CVs of 24-33% for cores

collected in the microcosms. Only for the cores collected in situ, the CV was relatively

high with 42%.

Discussion

Intracellular nitrate in intertidal sediment

A large pool of intracellular nitrate was discovered in the sediment of an intertidal flat

in the German Wadden Sea. Intracellular nitrate exceeded porewater nitrate levels and

was also present at depths where porewater nitrate was depleted. The high abundance of

diatoms in this and other intertidal flats (MacIntyre et al. 1996) suggests that nitrate is

stored in benthic phototrophic microorganisms. Correlative evidence for nitrate storage

in diatoms was obtained by photopigment analysis in the intertidal sediment incubated

in laboratory microcosms. Sedimentary pools of intracellular nitrate due to nitrate

storage by diatoms may be a wide-spread phenomenon in coastal marine sediments,

since diatoms dominate microphytobenthic communities in intertidal flats (MacIntyre et

al. 1996) and are able to store nitrate intracellularly (Garcia-Robledo et al. 2010, Kamp

et al. 2011). Nevertheless, the presence of intracellular nitrate might also be linked to

large sulphur bacteria (Sayama 2001), but in the sulphide-poor intertidal sediment near

172


Dorum-Neufeld (Jahn & Theede 1997), these microorganisms were not present, as

confirmed by microscopy. Also many benthic foraminifera are able to store nitrate, but

obviously not the species occurring in German Wadden Sea sediments (Risgaard-

Petersen et al. 2006, Pina-Ochoa et al. 2010).

The maximum concentration of intracellular nitrate was found in the upper 5 cm of the

intertidal sediment, which contrasts with the thin layer of intracellular nitrate at the

surface of other coastal marine sediments (Lomstein et al. 1990, Garcia-Robledo et al.

2010). Interestingly, the layer of high intracellular nitrate concentrations is often much

thicker than the photosynthetically active layer (Fenchel & Straarup 1971). This broad

distribution of intracellular nitrate must be caused by the presence of diatoms in deep,

aphotic sediment layers. The gliding motility of diatoms allows them to migrate

vertically in the sediment, but only down to depths of a few millimetres. In contrast,

passive burial of diatoms may relocate them several centimetres or decimetres into the

sediment. Known burial mechanisms of small microalgal cells are advective porewater

flow in permeable sandy sediments (Huettel & Rusch 2000, Ehrenhauss et al. 2004) and

bioturbation by animals such as the polychaetes H. diversicolor and A. marina that were

abundant at the time of sampling the intertidal flat.

Effect of H. diversicolor on intracellular nitrate

In the laboratory microcosms, benthic diatoms were most probably the main nitrate-

storing organisms because intracellular nitrate concentrations correlated well with

distributions of chlorophyll a and fucoxanthin. Large sulphur bacteria and foraminifera

did probably not contribute substantially to the sedimentary pool of intracellular nitrate

for the reasons given above. The intracellular nitrate pool was larger in the Hediste than

in the Control microcosm, even though diatom density and distribution were the same in

the two treatments. This means that H. diversicolor increased the average concentration

of intracellular nitrate in the diatom cells rather than changing the cell density of nitrate-

storing diatoms. The presence of H. diversicolor also enhanced the sedimentary

nitrification rate and enlarged the zone in which nitrification took place. Nitrification

might be stimulated by the increased oxygen availability due to searching and foraging

activities of the polychaete. In deeper sediment layers, burrow ventilation and

ammonium excretion may increase both oxygen and ammonium availability and

173


consequently nitrification in the thin oxic layer of the worm burrows (Mayer et al. 1995,

Nielsen et al. 2004). However, the excretion activity of the polychaete may also affect

nitrification at the sediment surface, if a substantial fraction of the ammonium excreted

is expelled into the water column due to the worm’s ventilation activity. This effect

might be quantitatively important in shallow coastal ecosystems and reinforced in a

recirculating system as used here. In line with this reasoning, the Ammonium treatment,

meant to mimic the ammonium excretion by H. diversicolor (Christensen et al. 2000),

increased the porewater and intracellular nitrate concentrations close to the sediment

surface. The increased availability of ammonium alone was sufficient to enlarge the

nitrate pools, while the increased availability of oxygen seemed to be less important for

the stimulation of nitrification.

The microcosm experiment revealed that diatoms take up and store more nitrate

intracellularly when nitrifying bacteria produce nitrate in the immediate environment of

the algae, maybe because of the efficient transport of nitrate within the oxic sediment

layer. The tight relationship between sedimentary nitrification and the storage of

intracellular nitrate by diatoms was demonstrated by the addition of the nitrification

inhibitor ATU to the second Hediste treatment. No stimulatory effect of H. diversicolor

on the intracellular and porewater nitrate pools occurred in the presence of ATU, despite

the worms’ ammonium excretion and sediment oxygenation.

Also the flux of dissolved inorganic nitrogen (DIN) between the water column and the

sediment has probably affected the nitrate pools in the sediment. The DIN

concentrations in the water column that drove these fluxes differed between the four

treatments in an expected way. The North Sea water supplied to the microcosms

contained ca. 40 μmol L�1 nitrate and the average nitrate concentration remained close

to this concentration, except for the Hediste + ATU treatment where it decreased in the

water column. This observation was in line with nitrification being inhibited and nitrate

consumption going on in the sediment. Together, the missing nitrate production in the

sediment and the lower nitrate flux from the water column explain the small

sedimentary pool of intracellular nitrate observed in the ATU-treated sediment. The

North Sea water supplied to the microcosms contained ca. 1-2 μmol L�1 ammonium and

this concentration remained unchanged, except for the Ammonium treatment where it

was deliberately adjusted to a concentration higher than in natural North Sea water and

174


also in the Hediste + ATU treatment where it increased due to the worms’ ammonium

excretions and nitrification being inhibited in the sediment. The possible ammonium

flux from the water column into the sediment increased the intracellular nitrate pool

only in the Ammonium treatment, but not in the Hediste + ATU treatment, which

underlines that ammonium must be nitrified to nitrate in the sediment to exert a

measurable effect on the intracellular nitrate pool.

Extrapolation to natural conditions

After the equilibration time of 10 days, worm distribution in the sediment microcosms

was relatively homogenous, while the diatoms that were visible on the sediment surface

occurred in patches. The very thin tip of microsensors measures chemical gradients in a

very small spot and these gradients may differ considerably even between neighbouring

spots. To arrive at representative oxygen and nitrate gradients, microprofiles were

repeated at as many randomly chosen spots as possible in each microcosm (i.e., up to

14). In contrast, sediment cores with a diameter of 2.5 cm integrate the measured

parameter over an area of ca. 5 cm2. Replicate sediment cores were thus expected to be

more similar than replicate microprofiles and therefore coring was repeated only 4 times

in each microcosm. In fact, the observed within-treatment variability of depth-integrated

profiles was high for microsensor data, intermediate for intracellular nitrate data, and

low for pigment data. In all cases, the within-treatment variability was low enough to

allow for comparisons between the treatments, and also lower than the one observed

under in situ conditions.

The intracellular nitrate concentrations in the sediment microcosms were in the range of

naturally occurring concentrations in coastal marine sediments (Garcia-Robledo et al.

2010, Høgslund et al. 2010). The difference in the vertical distribution of intracellular

nitrate concentrations between the sediment microcosms and the field site are probably

due to differences in the flow regime (i.e., tidal currents vs. continuous flow) and the

community of burrowing macrofauna (i.e., single-species vs. multi-species community).

In the sediment microcosms, the bioturbation and bioirrigation activities of

H. diversicolor did not lead to substantial burial of diatoms. The density of

H. diversicolor in the sediment microcosms was in the lower range of densities reported

for marine sediments (Scaps 2002). The effect of H. diversicolor on intracellular nitrate

175


pools might thus be even stronger at higher worm densities. The nutrient concentrations

were in the range of naturally occurring concentrations in the Wadden Sea (Kieskamp et

al. 1991, van Beusekom et al. 2008). Chlorophyll a concentrations in the sediment

microcosms were in the normal range (MacIntyre et al. 1996). The observed effects of

H. diversicolor on the sedimentary pool of intracellular nitrate might thus be

representative for intertidal flats of similar sediment composition. It is noteworthy that

the quantitative importance of intracellular nitrate will be particularly high in low-

porosity sediments as studied here, whereas porewater nitrate is likely more important

in high-porosity sediments such as mud.

Possible fate of intracellular nitrate

When integrating the vertical profiles over depth, the intertidal sediment contained

619 μmol m�2 intracellular nitrate and 87 μmol m�2 porewater nitrate. Intracellular

nitrate may thus sustain nitrate-consuming processes in the sediment longer or at a

higher rate than porewater nitrate. The diatoms themselves use intracellular nitrate for

assimilation (Lomas & Glibert 2000), but have also been shown to reduce it to

ammonium in a dissimilatory process that is induced by dark, anoxic conditions (Kamp

et al. 2011). Both diatoms and intracellular nitrate have been detected in anoxic

sediment layers of the sediment microcosms and thus diatoms do not entirely consume

their intracellular nitrate content under dark, anoxic conditions. It may be speculated

that some nitrate leaks out of the diatom cells (e.g., upon lysis following the freezing of

sediment at low tide during winter) and fuel anaerobic nitrate respiration by other

microorganisms in the sediment. Denitrification rates reported for intertidal sediments

in the Wadden Sea range from 0.2 to 190 μmol N m�2 h�1 (Kieskamp et al. 1991, Jensen

et al. 1996, Gao et al. 2010). Hence, intracellular nitrate from diatoms might sustain

denitrification in intertidal sediments for 3-129 days. Both anaerobic and aerobic

denitrification occur in Wadden Sea sediments (Gao et al. 2010) and could be fuelled by

intracellular nitrate from diatoms that are present in both oxic and anoxic sediment

layers.

176


Conceptual model of macrofauna effect on intracellular nitrate

We propose a mechanism of nitrogen cycling in coastal marine sediments in which

organisms from different trophic levels interact in converting particulate organic

nitrogen (PON) to nitrate that is stored in sedimentary microorganisms (Fig. 6).

PON NH4+

NH4+

Anaerobic NO3-

respiration

Burial into anoxic layers

NO3- storage

NO3- assimilation

NO3-

Nitrification

Excretion

Ventilation

NO3-

Feeding

O2

Water column

Sediment

O2

PON NH4+

NH4+

Anaerobic NO3-

respiration

Burial into anoxic layers

NO3- storage

NO3- assimilation

NO3-

Nitrification

Excretion

Ventilation

NO3-

Feeding

O2

Water column

Sediment

O2

Figure 6: Conceptual model of nitrogen cycling in intertidal sediments as affected by nitrate-storing diatoms and burrowing polychaetes. Worms feed on organic matter (black circles) from the water column or the sediment surface, excrete ammonium and oxygenate the sediment by their foraging, burrowing, and ventilation activities. Thereby, worms enhance the activity of nitrifying bacteria (white circles) that oxidise ammonium to nitrate in oxic sediment layers. Nitrate is taken up and stored intracellularly by diatoms (perforated discs). The sedimentary pool of intracellular nitrate can be used for nitrogen assimilation or for anaerobic nitrate respiration in anoxic sediment layers.

Burrowing macrofauna feeds on organic matter and excretes ammonium or nitrogen-

rich organic compounds like mucus or silk. Additionally, they oxygenate the sediment

by their foraging, burrowing, and ventilation activities, which results in additional

interface area and enhanced solute fluxes. Macrofauna thereby stimulates the activity of

nitrifying bacteria that oxidise ammonium to nitrate in the oxic sediment layers. Nitrate

is then taken up by microalgae (e.g., diatoms) and stored intracellularly. This

177


mechanism leads to a change in the forms of nitrogen (nitrate vs. ammonium and PON)

and in the compartmentalisation of nitrogen (intracellular vs. extracellular). The

conversion of PON into reactive oxidised nitrogen may fuel benthic primary production

or serve as an electron acceptor in anoxic sediment layers.

It remains to be investigated whether macrofauna also influences the fate of intracellular

nitrate in coastal marine sediments. It may be speculated that macrofaunal activities will

determine whether nitrate-storing microorganisms use their intracellular nitrate

themselves or whether it is made available to other sediment microorganisms. Sediment

reworking and burrow construction by macrofauna may relocate microphytobenthos

within the sediment and expose it to different microenvironmental conditions (e.g., from

light to dark, or from oxic to anoxic conditions). In the light, the microalgae probably

use intracellular nitrate for nitrogen assimilation (Lomas & Glibert 2000). If buried by

macrofauna to dark and anoxic conditions, the microalgae may use their intracellular

nitrate for dissimilatory nitrate reduction to ammonium (DNRA) to meet the energy

demand for entering a resting stage for long-term survival (Kamp et al. 2011).

Conversely, macrofauna that feeds on microphytobenthos may cause the lysis of nitrate-

storing cells in the gut (Smith et al. 1996). Denitrifying bacteria in the gut can use

nitrate leaking out of the lysing microalgal cells and produce nitrous oxide and

dinitrogen gas which are emitted from the animal (Stief et al. 2009, Heisterkamp et al.

2010). Nitrate that is not used in the gut will be excreted and can be used by sediment

bacteria in the immediate surrounding of macrofauna burrows as possible hotspots of

nitrate respiration.

178


Acknowledgements

We are grateful to the technicians of the Max-Planck-Institute for Marine Microbiology

for the construction of microsensors. Markus Huettel is acknowledged for inspiring

discussions. This study was financially supported by a grant from the German Science

Foundation awarded to P.S. (STI 202/6) and by the Max-Planck-Society, Germany.

179


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Chapter 7

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Chapter 7 Conclusion and perspectives

Conclusion and perspectives

This thesis focuses on interactions between benthic aquatic invertebrates and microbes

and their role in biogeochemical nitrogen cycling with particular emphasis on the

production of the gas N2O. Although N2O is a very important greenhouse gas and

ozone-depleting substance, the global N2O budget remains poorly quantified, in

particular with respect to biogenic N2O sources (Forster et al. 2007). This thesis

presents the first results on N2O emission from marine macrofauna that has so far been

overlooked as a biogenic source of N2O. The dataset compiled here contributes to the

currently small record on N2O emission by invertebrate species and extends our

knowledge on animal-associated N2O production from terrestrial and freshwater

habitats to the marine realm. Furthermore, it complements earlier studies on N2O

emission from terrestrial and freshwater invertebrates by identifying microbial biofilms

on the external surface of invertebrates as site of intense N2O production and

nitrification as important pathway involved in animal-associated N2O production. In

conclusion, benthic invertebrates can stimulate microbial N2O production by providing

three distinct habitats with specific microenvironments:

a) the gut, a transient microbial habitat characterized by low O2, high Corg and NO3�

concentrations, which is thus favourable for denitrification (Drake et al. 2006, Stief

et al. 2009 Chapters 2, 4, 5),

b) the exoskeleton or shell surface, a relatively persistent habitat exposed to fluctuating

O2 concentrations and high inorganic N input that provides suitable conditions for

the formation of microbial biofilms, in which nitrification and denitrification can

(co-) occur (Chapters 2-4),

c) the burrow, a microbial habitat with fluctuating environmental conditions and steep

concentration gradients, in which nitrification and denitrification can prevail in close

proximity (Svensson 1998, Stief & Schramm 2010).

By creating these microsites, invertebrates influence microbial metabolism and alter

rates of nitrification and denitrification and their N2O yields and thus net production

rates of N2O. In the invertebrate-provided microsites, the net N2O production rates can

be far higher than in the surrounding environment, which can lead to high N2O emission

rates from soils, sediments and water bodies that are densely colonized by invertebrates

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(Drake & Horn 2007, Stief et al. 2009, Chapter 5). Benthic invertebrates thus represent

abundant hotspots of N2O production that need to be integrated conceptually and

quantitatively into the global N cycle.

Conceptual integration of N2O emission from aquatic invertebrates

N2O-emitting benthic coastal invertebrates were found among the taxonomic groups of

Crustacea, Mollusca, Polychaeta and Echinodermata that comprise most of the soft-

bottom macrofauna species in coastal marine habitats (Chapter 2). The ability to emit

N2O is thus widespread among benthic coastal invertebrates and appears to be the rule

rather than the exception. However, the potential N2O emission rate varies considerably

with species, depending on the body size, habitat, presence of microbial biofilms on

external surfaces, and conditions in the gut of the respective species (Chapters 2, 3, 5).

Microbial N2O production in the invertebrate gut

In the gut, N2O is only produced in significant amounts if high numbers of active

denitrifiers are present that produce more N2O than they consume. Consequently,

factors that influence N2O production in the gut are:

a) the feeding type and the feeding rate of the animal that determine the amount of

ingested bacteria,

b) the physico-chemical conditions (e.g., O2, Corg, NO3�, pH) in the gut that determine

the activity of denitrifiers and the balance of denitrification enzymes, hence the rate

and N2O yield of gut denitrification,

c) the gut residence time that determines how long ingested denitrifiers are exposed to

the specific conditions in the gut and thus how much time they have for expression

of the denitrification genes,

d) the lysozyme activity that determines the number of viable denitrifiers in the gut

(Horn et al. 2003, Stief et al. 2009, Stief & Schramm 2010, Chapters 2-5).

For freshwater invertebrates, the amount of ingested bacteria is an important factor,

since N2O emission rates depend strongly on the feeding type of the animal (Stief et al.

2009). This does not hold true for marine invertebrates, which indicates that the

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physico-chemical conditions, the gut residence time and/or the lysozyme activity are

more important in determining the net rate of N2O production in the gut.

In respect to O2, the gut microenvironment of most aquatic macrofauna can be assumed

to be conducive for denitrification. Dissected guts of the shrimp Litopenaeus vannamei

were hypoxic to anoxic depending on their filling state even when being exposed to

fully oxygenated medium and no oxygen gradients were observed along the gut axis

(Chapter 5). Diffusion of oxygen into the gut might only play a role in smaller species

that have smaller volume to surface ratios of the guts (Tang et al. 2011). Here, oxygen

gradients may prevail along the gut axis and/or radius due to diffusion of oxygen

through the gut wall or openings and anoxic conditions may only occur in some sections

or the core of the gut (Schmitt-Wagner & Brune 1999, Tang et al. 2011). However, even

in very small animals such as the insect larvae Chironomus plumosus the gut can be

completely anoxic if diffusion of oxygen from the haemolymph and uptake of oxic

water by feeding is not sufficient to oxygenate the gut contents (Stief & Eller 2006).

Moreover, it was shown for the freshwater mussel Dreissena polymorpha that both

nitrifiers and denitrifiers are present in the gut, but only the denitrifying bacteria are

metabolically active (Chapter 4). Overall, this indicates that the gut of aquatic

invertebrates constitutes a microenvironment of low oxygen concentration that supports

denitrification activity. However, so far only a limited number of studies characterized

the gut microenvironment of a few invertebrate species in respect to oxygen, nitrate,

organic carbon, pH, and other physico-chemical parameters, and further investigations

are needed to improve our knowledge about environmental conditions and associated

microbial metabolism in the guts of invertebrates.

Based on the few studies on the invertebrates’ gut microenvironments, it can be

expected that the physico-chemical conditions in respect to nitrate and organic carbon

are especially favourable for denitrification in the gut of aquatic detritivores such as

deposit- and filter-feeding invertebrates. These animals take up large amounts of

organic carbon and NO3� by ingesting water-soaked food particles (Stief et al. 2010).

Moreover, they might ingest large numbers of nitrate-accumulating diatoms that, after

being lysed, could supply additional nitrate for gut denitrification. Benthic diatoms from

an intertidal flat can store more NO3� intracellularly than is available in the porewater of

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the sediment (Chapter 6). Invertebrate species, like Hediste diversicolor, that enhance

the sedimentary pool of intracellular nitrate, might thus indirectly stimulate N2O

production by increasing the NO3� supply for gut denitrification in diatom-feeding

invertebrates.

However, even though physico-chemical conditions in the invertebrate gut are

favourable for denitrification, this does not necessarily lead to high net N2O production

rates. A high lysozyme activity, which is common among marine deposit- and filter-

feeding invertebrates, can lead to the digestion of a great fraction of the ingested

bacteria (McHenery et al. 1986, Plante & Shriver 1998), thereby reducing the amount of

actively denitrifying bacteria in the gut. Additionally, if invertebrate species have long

gut residence times and rather stable anoxic conditions in their guts, ingested denitrifiers

will have enough time to express the full set of denitrification genes and perform

complete denitrification. In the latter case, denitrification in the gut might prevail at

high rates, but produce only trace amounts of N2O.

In contrast, high N2O yields from gut denitrification can be expected for species with

short gut residence times because the induction of the N2O reductase lags behind that of

the other denitrifying enzymes and ingested denitrifiers are probably not long enough in

the anoxic gut to establish the complete denitrification pathway (Philippot et al. 2001,

Zumft & Körner 2007, Stief et al. 2009). Additionally, high N2O yields may arise from

oxygen and pH gradients along the alimentary tract because the N2O reductase is

efficiently inhibited by elevated oxygen concentration or low pH (Bonin & Raymond

1990, Drake & Horn 2007, Richardson et al. 2009). For the shrimp L. vannamei, for

instance, the very short gut residence time of maximal 1 h (Beseres et al. 2005) and the

constantly low oxygen and high pH conditions throughout the filled gut suggest that the

high N2O/N2 ratio from gut denitrification was due to delayed expression of the N2O

reductase (Chapter 5). In other species, however, oxygen and pH conditions might be

more important than gut residence time for high N2O yields from gut denitrification.

Notably, the gut of soil-feeding termites is highly structured and characterized by

changing pH and steep oxygen gradients across the gut radius and length (Brune &

Kuhl 1996, Schmitt-Wagner & Brune 1999). Furthermore, the nitrate and/or nitrite

concentrations in the invertebrate gut can exceed the concentrations in the ambient

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surrounding and might be high enough to inhibit the reduction of N2O (Blackmer &

Bremner 1978, Firestone et al. 1979, Drake & Horn 2007, Stief et al. 2009). The

reduction of N2O to N2 makes up only about 20% of the energy yield of denitrification

(Richardson et al. 2009) and reduction of NO3� and NO2

� might be preferred over

reduction of N2O when NOx� is in ample supply.

In conclusion, several environmental, autecological, and physiological factors may lead

to an imbalance in the activity of the different denitrification enzymes in the gut of

invertebrates and thereby to increased N2O yields from gut denitrification. Depending

on the invertebrate species and environmental conditions, the importance of each factor

may vary, which makes it difficult to predict N2O production by invertebrates. The

combination of environmental, autecological, and physiological controlling factors

makes denitrification in the gut of invertebrates different from denitrification in

sediments, soils, and water bodies, and a site of intense microbial N2O production.

Microbial N2O production in exoskeletal biofilms

Besides N2O production in the gut of invertebrates, microbial biofilms on hard external

surfaces of aquatic invertebrates were found to be sites of high N2O production. Studies

on three marine mollusc species from different habitats and feeding guilds and one

freshwater species revealed that shell biofilms contribute 18-96% to the total N2O

emission of the animals (Chapters 3 + 4). The widespread distribution and importance

of such biofilms are further supported by the significant positive correlation of the N2O

emission rates of 19 marine invertebrate species with the presence of microbial biofilms

on exoskeleton and shell surfaces (Chapter 2). These results challenge earlier studies on

N2O emission from freshwater invertebrates that ascribed N2O production exclusively

to microbial denitrification activity in the anoxic gut (Stief et al. 2009, Stief & Schramm

2010). Since these studies did not specifically test for N2O production in exoskeletal

biofilms, it can be speculated that also for biofilm-bearing freshwater invertebrates, N2O

production in exoskeletal biofilms is a common trait that significantly contributes to the

total N2O emission of the animal.

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While N2O production in the invertebrate gut is only linked to denitrification, N2O

production in shell biofilms can derive from both denitrification and nitrification

(Chapter 3). Accordingly, N2O is produced by two functional groups of microorganisms,

nitrifiers and denitrifiers, whose activity is regulated by different environmental factors

(Devol 2008, Ward 2008). Importantly, the involvement of nitrifying bacteria in the

production of N2O means that not only NO3� and NO2

� are precursors and drivers for

animal-associated N2O emission, but also NH4+. This is of particular importance in the

light of the high NH4+ excretion rates of many invertebrate species, as well as their

feeding and bioturbation activities that stimulate remineralisation and release of NH4+

from the sediment (Blackburn & Henriksen 1983, Aller et al. 2001, Chapters 3, 4, 6).

Benthic invertebrates thereby significantly increase the availability of NH4+, which is

often limiting in oxygenated sediments and water bodies (Canfield et al. 2005). By

enriching dissolved inorganic nitrogen (DIN) in their immediate surroundings,

invertebrates stimulate the growth of microbial biofilms on the external surfaces of their

body and consequently also N2O production (Chapter 3).

Furthermore, the investigated mollusc species were found to excrete more than enough

NH4+ to sustain the nitrification-derived N2O production in their shell biofilms.

Moreover, the NO3� produced by biofilm nitrification might support N2O production by

denitrification if nitrification and denitrification are tightly coupled in shell biofilms,

and may also supply NO3� for denitrification in the animal’s gut. A substantial part of

the animal’s N2O emission may therefore be fuelled by the animal’s excretion and is

independent from environmental DIN supply. In conclusion, high N2O emission rates

may not only occur in nutrient-rich ecosystems, but also in the nutrient-enriched micro-

environment of invertebrates living in otherwise nutrient-poor macro-environments.

The animal’s self-sustained N2O emission might also overcome the sometimes

counteracting effects of DIN availability and temperature that control the rates of N

conversions and N2O production throughout the seasonal cycle in temperate regions. In

summer, when temperatures are high, the water column concentrations of DIN are

typically low, limiting the rates of nitrification and denitrification and concomitant N2O

production in the sediment and water column (Jorgensen & Sorensen 1985, Jorgensen &

Sorensen 1988, Rysgaard et al. 1995). In winter, the opposite is the case, and

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temperature is the limiting factor. High rates of denitrification and N2O emission from

sediments therefore occur during spring and autumn when moderate nitrate

concentrations coincide with moderate temperatures (Jorgensen & Sorensen 1985). A

very similar seasonal pattern was observed for N2O emission from insect larvae whose

N2O emission derived solely from denitrification in the gut (Stief et al. 2010, Stief &

Schramm 2010). For invertebrate species with a significant fraction of the emitted N2O

being produced in shell biofilms, the seasonal variability of N2O emission rates might

be different. The density of invertebrates is generally higher in summer than in winter

and consequently high NH4+ excretion rates and high temperatures are likely to coincide

in the animals’ microenvironment in summer (Gardner et al. 1993, Rysgaard et al.

1995). This may lead to high invertebrate-associated N2O emission rates in summer. It

would therefore be essential to monitor the seasonal in situ N2O emission rates of

invertebrate species whose N2O emission derives mainly from the shell biofilm and

whose NH4+ excretion rates are high.

Apart from the DIN availability, oxygen is a key factor controlling the process rates of

nitrification and denitrification and their N2O yields (Goreau et al. 1980, Betlach &

Tiedje 1981). The oxygen distribution in shell biofilms is very heterogeneous and the

vertical concentration gradients vary with biofilm thickness and light conditions

(Chapter 3). Anoxic bottom layers only establish in thick shell biofilms under dark

conditions when oxygen is not produced by photosynthesis and respiration by the

biofilm community consumes all oxygen that diffuses into the biofilm from the

oxygenated environment. The community composition of the biofilm (heterotrophs

versus oxygenic phototrophs) and the light regime to which the microbial biofilm is

exposed in the environment are therefore important for determining the prevailing

oxygen concentrations in shell biofilms. Under low-light conditions, denitrification and

nitrification were equally important for N2O production (Chapter 3). It can be

speculated that the N2O emission rates from shell biofilms are generally highest under

low-light conditions because (i) denitrification can prevail in hypoxic to anoxic

microsites and (ii) nitrification will not be light-inhibited. Furthermore, N2O yields of

nitrification and denitrification are highest under hypoxic conditions (Goreau et al. 1980,

Bonin & Raymond 1990), which preferentially establish in shell biofilms under low-

light conditions.

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Relative N2O yields of nitrification and denitrification in shell biofilms measured under

oxic and anoxic conditions, respectively, were 3.7-13.4% and thus higher than N2O

yields normally observed in aquatic sediments and water columns (Seitzinger 1988,

Bange 2008). Under in situ conditions, frequent changes in oxygen concentration are

likely to occur in the shell biofilm due to the animal’s respiration, feeding and migration

behaviour and these fluctuations in oxygen concentration might further increase the

N2O yield. In an artificially grown biofilm, changes in O2 concentration caused high

transient accumulation of N2O (Schreiber et al. 2009). It can also be speculated that the

N2O yield from denitrification in shell biofilms is increased because of higher numbers

of aerobic denitrifiers. It has been suggested that aerobic denitrification especially takes

place in environments with frequently changing oxygen conditions and that aerobic

denitrifiers preferentially produce N2O (Frette et al. 1997, Patureau et al. 2000, Gao et al.

2010). Moreover, imbalanced enzyme activity of nitrifiers and denitrifiers due to

frequently changing conditions can result in the accumulation of the intermediate nitrite

(Stein 2011), which was shown to strongly increase the N2O production in shell

biofilms (Chapter 3).

In conclusion, aquatic invertebrates provide distinct microenvironments in their guts

and on their external surfaces that stimulate microbial N2O production. These micro-

environments complement the known sites of N2O production in aquatic habitats, such

as sediments and biofilms. Conceptually, invertebrate-associated N2O production stands

out due to its complex control by environmental, autecological, and physiological

factors. In respect to the global N cycle, N2O production associated with aquatic

invertebrates constitutes a link between reactive nitrogen (i.e. nitrate, nitrite, and

ammonium) in aquatic ecosystems and the potent greenhouse gas N2O in the

atmosphere. Due to heterogeneous distribution, abundance and composition of the

invertebrate communities, and seasonal changes in environmental drivers, it can be

expected that the invertebrate-derived N2O emission from benthic aquatic systems is

spatially and temporally very variable. The ecological concept of “hotspots” and “hot

moments” may thus be most appropriate to describe the N2O emission from

invertebrate-colonized benthic habitats.

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Quantitative integration of N2O emission from aquatic invertebrates

The high abundance and potential N2O emission rates of many aquatic invertebrates,

especially marine species, suggest that aquatic invertebrates significantly contribute to

the overall N2O emission from aquatic environments. However, estimations on in situ

rates of N2O emission from aquatic invertebrates are difficult due to high variability

between species and the multiple temporally changing factors that control invertebrate-

associated N2O production. Furthermore, interactions between the invertebrates and the

environment might reduce or further stimulate N2O emission depending on the exact

site at which the invertebrates emit N2O and the ambient N2O production and

consumption rates (Stief & Schramm 2010). At the current state of research, only a very

rough quantitative integration of invertebrate-associated N2O production can be made

based on potential N2O emission rates because in situ measurements throughout the

year are missing.

Generally, it can be expected that N2O emission from aquatic invertebrates is highest in

nutrient-rich environments like coastal areas, lakes and streams, since these

environments sustain high numbers of invertebrates due to high primary production and

supply plenty of inorganic and organic substrates (Beukema et al. 2002). The Wadden

Sea is a very productive system with usually dense populations of diverse epi- and

infaunal invertebrates (Beukema et al. 2002). A typical average density of macrobenthic

fauna in tidal flats of the Wadden Sea is around 2000 individuals m-2 (Beukema 2002).

Assuming that the intertidal macrobenthic fauna emit N2O at the average potential

emission rate of 0.32 nmol N2O individual-1 h-1 determined for 19 benthic coastal

invertebrate species, a population of 2000 individuals m-2 results in an areal N2O

emission rate of about 1.28 μmol N m-2 h-1 or about 11 mmol N m-2 per year. This is

within the range of -2.9 to 8.6 μmol N m-2 h-1 reported for N2O fluxes from intertidal

and estuarine sediments (Seitzinger 1988, Kieskamp et al. 1991, Middelburg 1995, Usui

et al. 2001), which probably already include the effect of the benthic invertebrate

community on N2O emission rates if measurements were done in macrofaunal-

colonized sediments. According to these calculations, N2O production associated with

invertebrates accounts for a significant fraction of N2O fluxes from coastal marine

sediments. Whether the average potential N2O emission rate of 0.32 nmol ind.-1 h-1

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reflects the average annual in situ N2O emission rate of the benthic invertebrate

community is, however, highly uncertain. It can be speculated that the average annual in

situ N2O emission rate of macrofauna from temperate coastal areas is lower than

0.32 nmol ind.-1 h-1, since throughout much of the year the in situ temperatures are

lower than the temperature at which the potential emission rates were measured. On the

other hand, invertebrate densities in shallow coastal habitats can be much higher than

2000 individuals m-2 from spring to autumn due to extremely high abundances of N2O-

emitting mollusc species like Hydrobia ulva and Macoma balthica (Beukema et al.

1996, Barnes 1999, Ysebaert et al. 2005).

Extrapolation of the estimated annual N2O emission from benthic invertebrates of

11 mmol N m-2 to the complete area of the Wadden Sea of 13 000 km2 (van Beusekom

& de Jonge 2002) results in the emission of about 0.002 Tg N yr-1 (Table 1). This is ca.

1% of the estimated global N2O emission rate of earthworms (Drake et al. 2006).

Considering that benthic invertebrates colonize a great proportion of the world’s

continental shelf areas at relative high abundance (Bolam et al. 2010, Laverock et al.

2011), the global N2O emission rates of marine invertebrates and terrestrial earthworms

may be in the same order of magnitude. Assuming that the continental shelf area of

approximately 29 × 106 km2 (Inman & Scott 2005) is colonized with a mean macro-

faunal density of 500 individuals m-2 that emit N2O at a mean rate of 0.1 nmol

individual-1 h-1, marine invertebrates from continental shelf sediments would cause N2O

emissions of 0.35 Tg N yr-1. This is about 6% of the global N2O emission of

5.5 Tg N yr-1 from aquatic ecosystems (Denman et al 2007).

Table 1: Estimates of annual global N2O emissions from various sources

Source of N2O emission Tg N yr-1 Reference

Benthic macrofauna Wadden Sea 0.002 This thesis

Benthic macrofauna continental shelves 0.35 This thesis

Earthworms 0.19 Drake et al. 2006

Aquacultured shrimp Litopenaeus vannamei 0.0001 This thesis

Aquatic ecosystems 5.5 Denman et al. 2007

Total global N2O emission 17.7 Denman et al. 2007

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How much of the N2O produced by benthic marine invertebrates in continental shelf

sediments will actually reach the atmosphere depends on the hydrodynamics, the

distance of the N2O production site to the atmosphere and microbial N2O consumption

in the sediment and water column (Meyer et al. 2008, Bange 2008). The contribution of

N2O emission from marine invertebrates to the atmospheric N2O flux most likely

decreases with increasing water depth because more N2O can be consumed on the way

from the sediment to the atmosphere and the abundance of benthic macrofauna

decreases typically with increasing water depth (Wei et al. 2010). The impact of

epifaunal invertebrate species on N2O fluxes to the atmosphere might be greater than

that of infaunal species, since they emit N2O directly into the water column or in case of

intertidal species directly to the atmosphere, whereas N2O emitted by infaunal species

might be partly consumed in the surrounding sediment.

Conditions that favour N2O emission from aquatic invertebrates to the atmosphere

(i.e. high animal density, high nutrient concentrations, short distance between the N2O

production site and the atmosphere) are typically found in aquaculture facilities. The

shrimp L. vannamei is one of the most important aquaculture species that is reared at

high animal densities, temperatures, and nutrient concentrations. The high N2O

production in the gut of L. vannamei most likely contributed to the several-fold

supersaturation of N2O in the rearing water (Chapter 5), which probably caused high

N2O flux from the aquaculture to the atmosphere. With a global aquaculture production

of 2.72 × 106 tonnes in 2010 (FAO 2012), this species could have caused N2O emissions

from aquacultures of 0.13 Gg N yr-1, assuming that it emits N2O with an average rate of

0.2 nmol g-1 h-1 (Chapter 5). Since this N2O emission rate was measured under

conditions that were very similar to those normally found in aquacultures of

L. vannamei and conditions in the aquaculture farms are rather constant throughout the

year, this rate reflects reasonably well the annual in situ rate of N2O emission from L.

vannamei. The estimated global N2O emission from L. vannamei is rather insignificant

compared to the estimated total global N2O emission of 17.7 Tg N yr-1 (Denman et al.

2007). However, it is likely that also other aquacultured species (e.g., other shrimp and

prawn species, molluscs, and maybe also fish) emit N2O. Furthermore, it can be

expected that the contribution of aquacultured species to the global N2O emission will

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increase in the future due to the extremely fast-growing aquaculture industry (Williams

& Crutzen 2010).

N2O emissions from aquatic invertebrates in natural environments are also likely to

increase in the future due to the prospected increase in global warming and

anthropogenic N inputs to aquatic systems. The combination of higher temperatures and

increased availability of DIN will probably stimulate N2O emission from invertebrates

to the atmosphere, as these factors were shown to limit animal-associated N2O

production (Stief et al. 2010, Stief & Schramm 2010). Furthermore, moderate

eutrophication of aquatic systems leads to increased macrofaunal biomass and to

community shifts towards higher abundance of deposit- and filter-feeding species (Grall

& Chauvaud 2002, Nixon & Buckley 2002), which both are likely to enhance

invertebrate-associated N2O production.

Overall, the current dataset on N2O emission from aquatic invertebrates results in more

complete inventories of biogenic N2O sources, but does not reduce the uncertainties in

the global N2O budget. The first rough estimate on invertebrate-associated N2O

production suggests that aquatic invertebrates will only significantly contribute to N2O

emission from aquatic ecosystems if many abundant species emit N2O at high rates

throughout the year.

Future research

To arrive at more reliable estimates of the contribution of macrofauna to N2O emission

from aquatic environments, it is essential to measure in situ emission rates at high

resolution throughout the year to account for the expected spatial and temporal

variability in invertebrate-associated N2O production. Field measurements of N2O

emission rates and correlation with abundance and community composition of

invertebrates in different aquatic ecosystems and during all seasons are thus required.

Simultaneous monitoring of environmental drivers (ammonium, nitrite, nitrate,

temperature, oxygen, organic carbon, pH) is needed to improve our knowledge on

controlling factors of invertebrate-associated N2O production and their interaction with

each other.

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Since temperature and nitrate concentration have already been shown to be key factors

in regulating the N2O emission rate of aquatic invertebrates in temperate regions (Stief

et al. 2010, Stief & Schramm 2010), future studies should address for the first time

aquatic invertebrates in tropical regions. It can be assumed that the year-round high

temperatures and increasing eutrophication in tropical areas due to intensified

agriculture and aquaculture (Galloway et al. 2008) leads to exceptionally high

invertebrate-associated N2O emission rates in these regions. In the same context, it is

important to investigate the N2O emission potential of aquaculture key species that are

grown in very high numbers under extremely nutrient-rich conditions and at high

temperatures (FAO 2010). Both the per capita rates and the global rates of N2O

emission by aquacultured species may thus be far higher than from invertebrates in

natural environments.

Furthermore, it is essential to investigate the N2O emission potential of pelagic

invertebrate species. If also marine pelagic species emit N2O at significant rates, this

would increase the importance of invertebrate-associated N2O emission tremendously.

This is on the one hand due to the vast numbers of pelagic invertebrates in the world’s

oceans (e.g., krill, salps, copepods) and on the other hand due to direct emission of N2O

into the water column. Even if N2O emission rates from pelagic species are lower than

from benthic invertebrates, a higher fraction of the N2O emitted by pelagic invertebrates

may end up in the atmosphere due to lower N2O consumption rates in the water column

than in the sediment.

Special attention should be given to highly abundant aquatic invertebrate species. The

Antarctic krill Euphausia superba is estimated to be the species with the highest

biomass on Earth (379 million tonnes, Atkinson et al. 2009). The gut of Antarctic krill

is most likely a nitrate-rich microsite because this species inhabits the nitrate-rich

Southern Ocean (Kamykowski & Zentara 2005) and feeds primarily on diatoms that

have been found to accumulate nitrate intracellularly (Lomas & Glibert 2000, Kamp et

al. 2011). If this species emits N2O at a significant rate, krill alone could account for a

substantial fraction of oceanic N2O emission. Therefore, the next step will be to

investigate the rate and mechanisms of microbial N2O production associated with this

196


pelagic key species and examine whether the intracellular nitrate from the ingested

diatom cells can fuel denitrification in the krill’s gut.

Many open questions remain concerning the interaction between the host and the

associated microbes. I propose to specifically follow the fate of ammonium excreted by

the animals and quantify the amount that is metabolized to N2O in shell biofilms.

Furthermore, the animal may not only supply ammonium to the shell biofilm, but also

labile organic carbon compounds. Filter-feeding invertebrates generate a flow of

organic solutes and particles and motile invertebrates can actively search for food

sources, which as a side effect probably supply plenty of organic carbon to the shell

biofilm. The role of electron donors for invertebrate-associated N2O production has so

far not been studied and awaits further investigations. It is also not clear whether the

host is directly affected by the presence and activity of N2O-producing bacteria. Since

N2O is not toxic, the production in its body or on its external surface probably does not

harm the animal. In contrast, the animal may benefit from the presence of microbial

biofilms due to camouflage that reduces the predation pressure (Wahl 1989). Ingested

heterotrophic microbes, including denitrifiers, may support degradation of organic

matter in the animal gut and supply nutrients to the host or may compete with the host

for limited nutrients.

A combination of field measurements, controlled laboratory experiments, and molecular

analysis of the microbial communities is needed to investigate the complexity and

heterogeneity of processes and factors involved in N2O emission from aquatic

invertebrates. This is required to resolve the mechanisms, importance of regulating

factors, and eventually the contribution of invertebrate-associated N2O production to the

overall N2O emissions from aquatic environments.

197


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Contributed works

Contributed works

Denitrification in human dental plaque

Microscopic oxygen imaging based on

fluorescein bleaching efficiency measurements

203

Contributed works

Denitrification in human dental plaque

Frank Schreiber1, Peter Stief1, Armin Gieseke1, Ines M Heisterkamp1, Willy

Verstraete2, Dirk de Beer1, Paul Stoodley3,4


Celsiusstraße 1, 28359 Bremen, Germany.

2Laboratory of Microbial Ecology and Technology (LabMET), Ghent University,

Coupure Links 653, B-9000 Ghent, Belgium.

3Center for Genomic Sciences, Allegheny General Hospital/Allegheny-Singer Research

Institute, 320 East North Avenue, Pittsburgh 15212-4772,

Pennsylvania, USA.

4National Centre for Advanced Tribology at Southampton (nCATS), School of

Engineering Sciences, University of Southampton,

Highfield, Southampton SO17 1BJ, UK.

Published in BMC Biology 8:24, 2010

I.M. Heisterkamp performed the molecular analysis of dental plaque with help of A.

Gieseke and P. Stief and contributed to measurements of oral N2O emissions.

204

Contributed works

Abstract

Background: Microbial denitrification is not considered important in human-associated

microbial communities. Accordingly, metabolic investigations of the microbial biofilm

communities of human dental plaque have focused on aerobic respiration and acid

fermentation of carbohydrates, even though it is known that the oral habitat is

constantly exposed to nitrate (NO3�) concentrations in the millimolar range and that

dental plaque houses bacteria that can reduce this NO3� to nitrite (NO2

�).

Results: We show that dental plaque mediates denitrification of NO3� to nitric oxide

(NO), nitrous oxide (N2O), and dinitrogen (N2) using microsensor measurements, 15N

isotopic labelling and molecular detection of denitrification genes. In vivo N2O

accumulation rates in the mouth depended on the presence of dental plaque and on

salivary NO3� concentrations. NO and N2O production by denitrification occurred under

aerobic conditions and was regulated by plaque pH.

Conclusions: Increases of NO concentrations were in the range of effective

concentrations for NO signalling to human host cells and, thus, may locally affect blood

flow, signalling between nerves and inflammatory processes in the gum. This is

specifically significant for the understanding of periodontal diseases, where NO has

been shown to play a key role, but where gingival cells are believed to be the only

source of NO. More generally, this study establishes denitrification by human-

associated microbial communities as a significant metabolic pathway which, due to

concurrent NO formation, provides a basis for symbiotic interactions.

Table 1: Denitrification genes in dental biofilms of five volunteers

Volunteer NO3� reductase NO2

� reductase NO reductase N2O reductase

narG nirS nirK cnorB qnorB nosZ

A + + + - + +

B + + + - + +

C + + + - + +

D + - + - + +

E + NA NA - + NA Results are based on detection of PCR product with the expected size or on additional analysis of the sequence of the PCR product. NA = not analysed.

205

Contributed works

Figure 4: N2O formation in the human mouth is dependent on salivary NO2

�/NO3�

concentrations and the presence of dental plaque. (a) Correlation of oral N2O production and salivary NO2

�/NO3� concentration in 15 volunteers with unbrushed teeth. Each data point

represents the rate of oral N2O accumulation of one individual on a certain day (black circles). Some volunteers were sampled on more than 1 day resulting in 19 data points in total. Four volunteers were additionally sampled before and after drinking NO3

�-rich beetroot juice to increase salivary NO2

�/NO3� concentration and oral N2O accumulation (white circles connected

by dotted line). (b) Effect of oral hygiene on N2O accumulation rate in the mouth. Oral N2O accumulation rate of individuals before tooth brushing plotted against the N2O accumulation rate after tooth brushing (closed circles). In six individuals an antiseptic mouth rinse that affects bacteria in the entire oral cavity was applied after tooth brushing (open circles, each of the six individuals is represented by a unique colour). For example, an individual (dark green) with an oral N2O accumulation rate of 500 nmol/h reduced the rate to 290 nmol/h by tooth brushing. Subsequent application of a mouth rinse resulted in a rate of 110 nmol/h. The dashed line corresponds to the absence of an effect of oral hygiene on the oral N2O accumulation. The error bars indicate the standard error of five replicate measurements of the oral N2O accumulation rate.

206

Contributed works

Microscopic oxygen imaging based on fluorescein bleaching efficiency measurements

Martin Beutler1,2, Ines M. Heisterkamp1, Peter Stief1, Dirk de Beer1



2bionsys GmbH, Fahrenheitstr. 1, 28359 Bremen, Germany

Manuscript in preparation

I.M. Heisterkamp contributed to microsensor measurements and performed oxygen

measurements in shell biofilms of different marine mollusc species.

207

Contributed works

Abstract

Photobleaching of the fluorophore fluorescein in an aqueous solution is dependent of

the apparent oxygen concentration. Therefore, the analysis of the time-dependent

bleaching behaviour is a useful measure of dissolved oxygen concentrations that can be

combined with epi-fluorescence microscopy. The molecular states of the fluorophore

can be expressed by a three-state energy model. This leads to a set of differential

equations which describe the photobleaching behaviour of fluorescein. The numerical

solution of these equations shows that in a conventional wide-field fluorescence

microscope, the fluorescence of fluorescein will fade out faster at low than at high

oxygen concentration. Further simulation showed that a simple ratio function of

different time-points during a fluorescence decay recorded during photobleaching could

be used to describe oxygen concentrations in an aqueous solution. Additional findings

were that a careful choice of dye concentration or excitation light intensity could help to

increase sensitivity in the oxygen concentration range of interest. In the simulations, the

estimation of oxygen concentration by the ratio function was very little affected by the

pH value in the range of pH 6.5 to 8.5. Filming the fluorescence decay by a charge-

coupled-device (ccd) camera mounted on a fluorescence microscope allowed a

pixelwise estimation of the ratio function in a microscopic image. Use of a microsensor

and oxygen-consuming bacteria in a sample chamber enabled the calibration of the

system for quantification of absolute oxygen concentrations. Finally, the method was

employed to nitrifying biofilms growing on snail and mussel shells to estimate apparent

oxygen concentrations.

208

Contributed works

No inhibitor

Inhibitor of nitrification (ATU)

sealing

cover slip

objective lens

biofilm

shell

fluorescein

microscopic slide

No inhibitor

Inhibitor of nitrification (ATU)

sealing

cover slip

objective lens

biofilm

shell

fluorescein

microscopic slide

sealing

cover slip

objective lens

biofilm

shell

fluorescein

microscopic slide

Figure 8: Oxygen measurements in microbial biofilms on the shell surfaces of marine invertebrates based on fluorescein bleaching measurements. Top left: fluorescence microscope with camera by which time series of fluorescein bleaching were measured. Top right: drawing of measuring chamber with incubated biofilm-covered shell. Bottom left: reflectance images of the biofilms on the shell of the mussel Mytilus edulis. Bottom right: oxygen images after biofilm-covered shells of M. edulis were incubated in artificial seawater without inhibitor and with the nitrification inhibitor allylthiourea (ATU) for 5 minutes. Blue means oxygen-depleted, red oxygen-saturated.

209

Contributed works

Hinia�reticulata

0

100

200

300

ASW ATU DCMU

Oxy

gen�

(μm

ol�L

�1)

Mytilus�edulis

0

100

200

300

ASW ATU DCMU

Oxy

gen�

(μm

ol�L

�1)

Littorina�littorea

0

100

200

300

ASW ATU DCMU

Oxy

gen�

(μm

ol�L

�1)

Figure 9: Oxygen concentrations in biofilms on the shells of the three mollusc species Hinia reticulata, Mytilus edulis, and Littorina littorea. Intact shell biofilms were incubated in artificial seawater without inhibitor (ASW), with the nitrification inhibitor allylthiourea (ATU), and with the photosynthesis inhibitor 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU). After 5 minutes of incubation, oxygen concentration was measured by photobleaching.

210

Danksagung

Danksagung

Zuerst möchte ich mich herzlichst bei Prof. Dr. Bo Barker Jørgensen bedanken, dass er

mich als seine letzte Doktorandin am Max-Planck-Institut angenommen hat und für das

Verfassen des Erstgutachtens. Vielen Dank geht auch an Herrn Prof. Dr. Ulrich Fischer,

der so freundlich war das Zweitgutachten zu übernehmen. Herzlich bedanken möchte

ich mich ebenfalls bei den anderen Mitglieder des Prüfungskomitees Herrn Prof. Dr.

Victor Smetacek, Dr. Peter Stief, Kristina Stemmer und Johanna Wiedling.

Mein größter Dank gilt Peter Stief, meinem Betreuer und Mentor während meiner

Doktorarbeitszeit. Peter, ich habe dir so vieles zu verdanken und möchte mich von

ganzem Herzen für deine großartige Unterstützung in den letzten Jahren bedanken. Ich

habe deine humorvolle Art, dein stets offenes Ohr, deine Ideen und Ratschläge für die

Laborarbeit als auch beim Verfassen der Manuskript sehr geschätzt.

Großer Dank gilt natürlich auch Dirk de Beer, der meine Arbeit in der

Mikrosensorgruppe immer unterstützt hat und mir mit seinen Kommentaren und Ideen

weitergeholfen hat.

Einen ganz besonderen Dank möchte ich den lieben „Dänen“ aussprechen, Andreas

Schramm, Nanna Svenningsen, Lone Heimann, Maria Sigby-Clausen und Lars Peter

Nielsen für die sehr gute langjährige Zusammenarbeit, die ich sehr genossen habe. Ich

möchte mich an dieser Stelle auch bei der gesamten Mikrobiologiegruppe der

Universität Aarhus bedanken, die mich immer herzlich aufgenommen hat bei meinen

zahlreichen Besuchen und mir eine tolle Zeit im schönen Aarhus bereitet hat.

Vielen herzlichen Dank für die gute Zusammenarbeit gilt auch Anja Kamp, für die

vielen guten Gespräche und schönen gemeinsamen Stunden im Labor und in der

Teeküche, Gaute Lavik dafür, dass er mir die Geheimnisse der Massenspektrometrie

näher gebracht hat und für anregende Diskussionen, Frank Schreiber für seine

interessanten Anregungen und einer spannenden Arbeit, bei der wir selbst Probanden

waren und Martin Beutler, für die zahlreichen Stunden mit ihm vor dem Mikroskop und

guten Diskussionen.

211

Danksagung

Vielen Dank gilt natürlich auch der Mikrosensorgruppe, in der ich mich all die Jahre so

wohl gefühlt habe. Vielen Dank an die TA’s für ihre Hilfe, wann immer ich sie

gebraucht habe. Und an euch, liebe anderen Doktoranden der Mikrosensorgruppe, für

die motivierenden Gespräche, Diskussion über die Wissenschaft und das Leben. Haltet

durch, ihr habt es auch bald geschafft.

Ganz lieben Dank auch an meine „Mädels“ (Anna, Anne, Frauke, Julia, Kirsten, Sandra,

Ulli und Verena) für die vielen amüsanten Mittagspausen und Mädelsabende. Mein

besonderer Dank geht dabei an Anna und Anne, die immer für mich da waren und mich

aufgemuntert haben, wenn es mal nicht so gut lief.

Natürlich möchte ich auch meine Bürokollegen danken, Mohammad Al-Najjar, Kristina

Stemmer und Stefan Häusler, sowie allen vorherigen Bürokollegen. Vielen Dank für die

tolle Zeit und super Atmosphere.

Ganz herzlich möchte ich meinen Mitbewohnern Ela, Jutta, Sebastian und Mika danken.

Ich habe die gemeinsame Zeit mit euch unglaublich genossen und danke euch sehr, für

die gemütlichen Abende in der Küche und dafür, dass ihr mir ein so schönes Zuhause

bereitet habt. Ela, dir möchte ich besonders danken für deine Hilfe und dein Verständnis

vor allem in der Endphase der Arbeit.

Danke auch an alle meine Freunde, die in den letzten Jahren oft von mir hören mussten,

dass ich keine Zeit habe, aber immer verständnisvoll waren und mir viel Mut zu

gesprochen haben.

Zum Schluss möchte ich von ganzem Herzen meiner Familie danken; für ihre Liebe,

Vertrauen und Glauben an mich, der mir geholfen hat, die manchmal doch sehr

schwierige Zeit gut zu überstehen. Ohne euch hätte ich das nie geschafft. Ganz

besonders möchte ich dir danken, Matthias, für dein immerwährendes Verständnis,

deine Unterstützung und dass du immer für mich da bist.

212

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